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COPYRIGHT DEPOSIT; 



METHODS IN PLANT HISTOLOGY 



METHODS 



IN 



PLANT HISTOLOGY 



BY 

CHARLES J. CHAMBERLAIN, Ph.D. 

Instructor in Botany in the University of Chicago 



5 J ) -> -> 

0) 3D 



c . ' 



CHICAGO 

Gbe TIlniversltB of Cbicaao IPtess 

1901 



THE LIBRARY OF 
CONGRESS, 

Two Copies Received 

JUN. 27 1901 

Copyright entry 
GLASS ^-OtXc, N* 

/ /9&$ 

COPY B. 



COPYRIGHT, I9OI 

BY THE UNIVERSITY OF CHICAGO 

CHICAGO, ILLINOIS 



CK 












PREFACE. 

This book has grown out of a course in histological tech- 
nique conducted by the author at the University of Chicago. 
The course has also been taken by non-resident students through 
the Extension Division of the University. The Methods were 
published over a year ago as a series of articles in the Journal 
of Applied Microscopy ', and have called out numerous letters of 
commendation, criticism, suggestion, and inquiry. The work 
has been thoroughly revised and enlarged by about one-half. It 
is hoped that the criticism and suggestion, and also the expe- 
rience gained by contact with both resident and non-resident 
students, have made the directions so definite that they may be 
followed, not only by those who work in a class under the 
supervision of an instructor, but also by those who must work 
in their own homes without any such assistance. 

More space has been devoted to the paraffin method than to 
any other, because it has proved to be better adapted to the 
needs of the botanist. The celloidin method, the glycerine 
method, and free-hand sectioning are also described, and their 
advantages and disadvantages are pointed out. 

The first part of the book deals with the principles of fixing 
and staining, .and the various other processes of microtechnique, 
while in the later chapters these principles are applied to spe- 
cific cases. This occasions some repetition, but the mere pres- 
entation of general principles will not enable the beginner to 
make good mounts. 

The illustrations and notes in the later chapters are not 
intended to afford a study of general morphology, but they 



vi Methods in Plant Histology 

merely indicate to students with a limited knowledge of plant 
structures the principal features which the preparations should 
show. The photomicrographs were made from the author's 
preparations by Dr. W. H. Knap, and figs. 52, 57, and 59 were 
drawn by Miss Eleanor Tarrant ; all other figures of plant 
structures were made from the author's drawings. 

Corrections and suggestions will be heartily appreciated. 

Charles J. Chamberlain. 
Chicago, 
June 1, 1901. 



CONTENTS. 

PART I. 

Page 

Chapter I. Apparatus i 

Chapter II. Reagents 7 

Killing and Fixing Agents 7 

Stains 7 

Formulae for Alcohols 9 

Miscellaneous Reagents 10 

Arrangement of the Outfit 10 

Chapter III. Temporary Mounts 11 

Chapter IV. The General Method 13 

Killing and Fixing 13 

Washing 14 

Hardening and Dehydrating 14 

Clearing 16 

The Transfer from Clearing Agent to Paraffin 16 

The Paraffin Bath 17 

Imbedding 18 

Cutting 19 

Fixing Sections to the Slide 20 

Removal of Paraffin 21 

Removal of Xylol 21 

The Transfer to the Stain 21 

Clearing 22 

Mounting in Balsam 22 

A Tentative Schedule for Paraffin Sections 23 

Chapter V. Killing and Fixing Agents 25 

The Alcohols 25 

The Chromic Acid Group 26 

Picric Acid 30 

Corrosive Sublimate 30 

Formalin 31 

General Hints on Fixing 31 

Chapter VI. Staining 33 

The Haematoxylins 34 

The Carmines 39 

The Anilins 41 

vii 



viii Methods in Plant Histology 

Chapter VII. General Remarks on Staining 47 

Chapter VIII. Practical Hints on Staining 53 

Chapter IX. The Celloidin Method 55 

Chapter X. The Glycerine Method 58 



PART II. 

Chapter XI. Thallophytes — Alg^e . . 63 

Cyanophycese . 63 

Chlorophyceae . , 65 

Phaeophycese 73 

Rhodophycese ■ 75 

Chapter XII. Thallophytes — Fungi 77 

Schizomycetes 77 

Myxomycetes 78 

Phycomycetes "."'-. . . . . .... . . 79 

Ascomycetes ' 80 

^Ecidiomycetes 84 

Basidiomycetes 87 

Lichens 88 

Chapter XIII. Bryophytes — Hepatice 89 

Chapter XIV. Bryophytes — Musci 97 

Chapter XV. Pteridophytes — Filicine^e 103 

Chapter XVI. Pteridophytes — Equisetine^e 115 

Chapter XVII. Pteridophytes — Lycopodine^e 117 

Chapter XVIII. Spermatophytes— Gymnosperms 119 

Chapter XIX. Spermatophytes — Angiosperms 129 

Chapter XX. Labeling and Cataloguing Preparations ... 141 

Chapter XXI. A Class List of Preparations 143 

Chapter XXII. Formulae for Reagents . 149 

Index 157 



Part I. 



CHAPTER I. 



APPARATUS. 

The following list of apparatus includes a fair equipment for 
histological work : a microscope magnifying at least 400 diam- 
eters ; a hand microtome; a sliding microtome ; a razor; a hone 
and a good razor strop ; a paraffin bath and lamp ; a turn-table ; 
a scalpel ; a pair of needles ; a pair of scissors ; a pair of for- 
ceps ; stender dishes ; minots or watch-glasses ; a wash bottle ; 
a graduate (50 or 100 cc); pipettes; slides, 1X3 inches; round 
covers, 18 mm. or ^ inch in diameter; and 
square covers, 7/q inch. Long covers, 22 X 
50 mm., will be needed for some of the serial 
sections. 

A convenient and effective microscope 
should have a rack and pinion coarse adjust- 
ment, a fine adjustment, two eyepieces (about 
one-inch and two-inch preferred), a low-power 
objective of two-thirds of an inch or a one-inch 
focus, a high-power objective of one-fifth or 
one-sixth of an inch focus, a double nose- 
piece, an iris diaphragm, and an Abbe con- 
denser. A cheap and practical form is shown 
in fig. 1, and similar instruments are for sale 
by all the leading companies. 

The hand microtome {fig. 2) will be found 

1 c , -111,11 . i of hand microtome. 

extremely useful, especially by the busy teacher 
who has large classes. Any sliding microtome, if kept in good 
order, will be sufficient for the work to be described in this book, 
but those of medium size are to be preferred. The student's 
microtome {figs, 3 and 3 A) is quite inexpensive and does good 




Methods in Plant Histology 




Fig. i. A compound microscope, with rack and pinion coarse adjustment, micrometer screw fine 
adjustment, triple nosepiece, iris diaphragm, and condenser. 



Apparatus 




WOrk Where eX- F IG «3- The student's microtome 

pense is not an 
objection the Jung c-~"~~" 
Thoma, the Minot, 
and other micro- 
tomes of similar 
grade are to be 
preferred. 

The stout ra- 
zors our grand- 
fathers used to 
shave with are ex- 
cellent for free- 
hand sectioning, 
for hand-micro- %V -C s -C****«^ T " 1" 

tome work, and ~-~ --,:"" 

even iOr Cutting FlG 3 A _ Clamp to hold an ordinary razor in the student's microtome. 




I 



Methods i?i Plant Histology 




paraffin sections on the sliding microtome. The blade should 
be flat on one side {fig. 4 A). Modern razors {fig. 4 B) with 
delicate blades, though good to shave with, are worthless for 
cutting sections of plants. The razor is a necessity; if a micro- 
tome knife is 
wanted in addi- 
tion, it should 
have a bevel 
about like that 
shown in fig. 4 A. 
FlG -4- Ashortblade,two 

or three inches in length, is to be 
preferred to the longer ones, which 
are much more troublesome to 
sharpen. 

There are numerous forms of 
the paraffin bath. Those with a 
water-jacket, a thermometer, and a 
thermostat to maintain an even 
temperature are the most con- 
venient where gas is available 




A paraffin bath, with water-jacket, 
designed to be used with a thermostat. 




A. 



B. 




r 



*« 



Fig. 6. A, top view; B, side view; C, end view; Z>, box to con- 
tain the paraffin. 



{fig. 5) . As a rule it 
is easier to keep the 
temperature constant 
in the larger baths. 
A bath which, if care- 
fully watched, gives 
the very best results 
can be made by any 
tinner, and is very in- 
expensive. The ac- 
companying figures 
show the form and 
dimensions {fig. 6). 

It is made of cop- 
per one thirty-second 



Apparatus 




Fig. 7. A, Naples jar; B, stender dish. 



of an inch thick, but thicker copper is as good or better. There 
should be two boxes to contain the paraffin ; the covers to the 
boxes should fit loosely. Any kind of a lamp may be used. 

Stender dishes are now very generally used for staining on 
the slide. The form shown in fig. 7 A is made just large enough 
to hold two slides, placed 
back to back, and hence 
requires only a minimum of 
the reagent. The cap in 
this form does not fit closely 
enough to keep absolute 
alcohol and xylol, but does 
very well for the other alco- 
hols and stains. We do not 
believe that the convex 
cover is as good as a flat 
one. The form shown in fig. 7 B is the best for absolute alco- 
hol and xylol, but even with this it is better to put a little 
vaseline or glycerine on the cover to prevent any evaporation. 

Wide-mouthed 
bottles, though 
not so conveni- 
ent, give just as 
good results. 

A serviceable 
form of turn-table 
for glycerine 
mounts is shown 
in fig. 8. 

The other 
pieces of appa- 
ratus mentioned 
need no com- 
ment. By consulting a catalogue, which will be furnished by any 
dealer, the beginner can determine what he needs to buy, and what 
he can find substitutes for, if it is necessary to be very economical. 




Fig. 8. Turn table. 



CHAPTER II. 

REAGENTS, 

It would require entirely too much space even to enumerate 
the reagents which are occasionally used in a fully equipped 
university laboratory. The following list includes only those 
which are used constantly. The Microtomisf s Vade-Mecum by 
Lee contains very complete formulae for stains and other reagents. 
The quantities mentioned below indicate about what the average 
student uses in a three-months' course in methods. Nearly all 
the stains, however, would last for a year, if properly used. 

KILLING AND FIXING AGENTS. 

Commercial alcohol (about 95 per cent.), 2 liters; absolute 

alcohol, 300 cc; ether, 50 cc; chromic acid, 10 g. ; corrosive 

sublimate, 10 g.; glacial acetic acid, 25 cc; hydrochloric acid, 

50 cc; picric acid, 5 g.; chloroform, 50 cc; [osmic acid, 1 per 

cent, solution in water, 25 cc. This is extremely expensive, and 

not necessary except for the most delicate work]. Formulae for 

making killing and fixing agents from these materials will be 

given later. 

STAINS. 

Only a few of the most important stains are given in this 
list. In general one should have enough of a stain to stand 
about two inches high in the stender dish or bottle in which the 
staining is to be done. The theory and practice of staining will 
be discussed later. 

Delafield's Hematoxylin. — To 100 cc. of a saturated solution of 
ammonia alum add, drop by drop, a solution of 1 g. haematoxylin 
dissolved in 6 cc. of absolute alcohol. Expose to air and light 
for one week, then filter. Add 25 cc. of glycerine and 25 cc. of 
methyl alcohol. Allow to stand until the color is rather dark. 
Filter, and keep in a tightly stoppered bottle. The solution 

7 



8 Methods in Plant Histology 

should stand for two months before it is ready for use, but, if 
needed immediately, the " ripening," which is brought about by 
the oxidation of haematoxylin into haematin, may be secured in 
a few minutes by a judicious addition of peroxide of hydrogen. 

Mayer's Haem-Alum. — Dissolve with gentle heat I g. of haema- 
toxylin in 50 cc. of 95 per cent, alcohol ; add a solution of 50 g. 
of alum in a liter of distilled water. Allow the mixture to cool 
and settle ; filter ; add a crystal of thymol to preserve from 
mold. The stain is ready for use as soon as made, and it keeps 
well. 

Haidenhain's Iron Alum-Haematoxylin.- — Two solutions are 
used, and they are never to be mixed : 

(a) A 1 ]/ 2 to 4 per cent, aqueous solution of ammonia sulphate of iron. 

(b) A y z per cent, aqueous solution of haematoxylin. 

Cyanin, Erythrosin, Safranin, Gentian Violet. — Numerous 
formulae are given for these and other anilin stains, but the fol- 
lowing general formula gives excellent results : 

Make a 3 per cent, solution of anilin oil in distilled water; 
shake thoroughly and frequently for a day; add enough alcohol 
to make the whole mixture about 20 per cent, alcohol. Add 1 g. 
of cyanin or erythrosin, etc., as the case may be, to 100 cc. of 
the solution. Safranin is often used in a strong alcoholic solu- 
tion, and even with the above formula it is better to dissolve the 
safranin in strong alcohol before adding it to the mixture. 

Acid Fuchsin. — Use a 1 to 2 per cent, solution in water, or 
70 per cent, alcohol. 

Iodine Green. — Use a 1 to 4 per cent, solution in water or 
alcohol. A 3 per cent, solution in 70 per cent, alcohol is very 
good for the vascular system of plants. 

Mixtures of Fuchsin and Iodine Green. — The following formula 
is often used for karyokinetic figures : 

(a) A y z per cent, solution of fuchsin in water. 

{p) A %. per cent, solution of iodine green in water. 

Just before using mix [a) and (£) in various proportions until 
what is needed for the particular case is found. 
Orange G. — Use a saturated aqueous solution. 



Reagents 



Eosin. — A I to 5 per cent, solution in water or alcohol. A 
2 per cent, aqueous solution is good for material to be mounted 
in glycerine, but a 2 per cent, solution in 70 per cent, alcohol is 
better for balsam mounts. The stronger solution maybe diluted 
as needed for special cases. 

FORMULA FOR ALCOHOLS. 

The grades of alcohol in most common use are 35 per cent., 
50 per cent., 70 per cent., 85 per cent., 95 per cent., and 100 per 
cent. The 100 per cent, is expensive, and great care should be 
taken to keep the bottle well corked or the stender dish closely 
covered. The following formulae will enable anyone to make 
the other grades of alcohol from 95 per cent, alcohol and water : 
95 35 95 50 95 70 95 85 



60 45 25 10 

The above are the formulae for 35 per cent., 50 per cent., 70 
per cent., and 85 per cent, alcohol. Any other grade can be 
gotten in the same way. In the first formula, subtract 35 from 
95 ; the result, 60, is the number of cubic centimeters of water 
which must be added to 35 cc. of 95 per cent, alcohol 
in order to obtain 35 per cent, alcohol. The mixture 
contains 95 cc. of 35 per cent, alcohol. If more or 
less than 95 cc. of the mixture is needed, take 
proportional parts of 35 and 60. This simple method 
is a time-saver, but if the bottles or stender dishes 
are to be filled frequently, it will be a still further 
saving of time to use a long label {fig. g) , and, after 
pouring in the 95 per cent, alcohol, draw a line 
showing how high it reaches, and then, after pouring 
in the water, draw another line. The next time it is 
necessary to fill the bottles merely pour in 95 per 
cent, alcohol until it reaches the first line, and then pour in water 
until it reaches the second line. It is not necessary to use dis- 
tilled water, if pure drinking water is available. 




Fig. 9. 



io Methods in Plant Histology 

CLEARING AGENTS. 

Xylol is the most generally useful clearing agent yet known. 
Clove oil, cedar oil, bergamot oil, carbolic acid, and turpentine 
are all necessary for special purposes. About 200 cc. of xylol, 
50 cc. of clove oil, and 25 cc. of each of the others makes a fair 
outfit to begin with. 

MISCELLANEOUS. 

Canada balsam, 25 cc; glycerine, 50 cc; glycerine jelly, 25 cc; 
2 per cent, celloidin, 50 cc; 10 per cent, celloidin, 50 cc; hard 
and soft paraffin, 500 g. each ; gold size, 25 cc; and a small soft 
brush for ringing glycerine mounts. 

ARRANGEMENT OF THE OUTFIT FOR STAINING AND MOUNTING. 

It is best to keep the various reagents in definite positions in 
order that no time may be lost in hunting for anything. The 




{ SaWmj UUW 



Fig. 10. 



accompanying diagram [fig. 10) of a part of the top of a table 
shows a convenient arrangement. 

The alcohols are in front and the stains are placed behind. 
The eosin, fuchsin, iodine green, cedar oil, and clove oil may be 
kept in bottles ; the rest should be in stender dishes. 



CHAPTER III. 

TEMPORARY MOUNTS. 

Before considering the complicated methods involved in 
making permanent preparations a word should be said about 
temporary mounts. A preliminary examination of almost any 
botanical material may be made without any fixing, imbedding, 
or staining. If a little starch be scraped from a potato, and a 
small drop of water and a cover-glass be added, a very good 
view will be obtained, and if a small drop of iodine solution be 
allowed to run under the cover, the preparation, while it lasts, 
could hardly be improved. The unicellular and filamentous 
algae can be studied quite satisfactorily from such mounts. 
The protonema of mosses and the prothallia of ferns should be 
studied in this way, even if a later study from sections is 
intended. If the top of a moss capsule be cut off at the level 
of the annulus, a beautiful view of the peristome may be obtained 
by simply mounting in a drop of water, or, in a case like this 
where no collapse is to be anticipated, the object may be mounted 
in a small drop of glycerine — just enough to come to the edge 
of the cover without oozing out beyond — and the preparation 
made permanent by sealing with gold size or any good cement. 
The antheridia and archegonia of mosses may be examined if 
the surrounding leaves are carefully teased away with needles. 
Free-hand sectioning with a sharp razor and judicious teasing 
with a pair of needles will give a fair insight into the anatomy 
of the higher plants without demanding any further knowledge 
of technique. This rough work is a very desirable antecedent to 
the study of microtome sections, because most students see in a 
series of microtome sections only a series of sections when, in 
the mind's eye, they ought to see the object building itself up in 
length, breadth, and thickness as they pass from one section to 
another. 



CHAPTER IV. 

THE GENERAL METHOD. 

We shall now consider the routine of mounting an object in 
Canada balsam. While the outline refers more particularly to 
the paraffin method, the principles are general in their applica- 
tion and must be mastered by everyone who desires to make 
first-class preparations. Several of the topics, like killing and 
fixing, staining, etc., will be treated in detail when considering 
the various reagents. 

I. KILLING AND FIXING. 

Usually the same reagent is used for both killing and 
fixing. The purpose of a killing agent is to bring the life- 
processes to a sudden termination, while a fixing agent is used 
to fix the cells and their contents in as nearly the living condi- 
tion as possible. The fixing consists in so hardening the 
material that the various elements may retain their natural con- 
dition during all the processes which are to follow. This step 
is one of extreme importance. Take the killing and fixing 
fluids into the field. If one waits until the material is brought 
to the laboratory, there may be some fixing, but it will, 
in many cases, be too late to do much killing. Material like 
Spirogyra, however, may be brought from the field into the 
laboratory before fixing, if considerable water be brought with 
it. Branches with developing buds may be brought in and kept 
in water. Always have the material in very small pieces, in 
order that the reagents may act quickly on all parts of the 
specimens. Pieces larger than one-fourth inch cubes should be 
avoided whenever possible. For very fine work no part of the 
specimen should require the reagent to penetrate more than 
one-sixteenth of an inch. In general, the volume of the reagent 
should be ten to fifty times that of the material. The time 

13 



14 Methods in Pla?it Histology 

required for this process varies with the reagent, the character 
of the tissue, and the size of the piece. About twenty-four 
hours is a commonly recommended period for chromic acid 
solutions. While this might suffice in some cases, we should 
recommend two or three days, and even a longer period would 
probably do no harm. 

II. WASHING. 

Nearly all fixing agents, except the alcohols, must be 
washed out from the material as completely as possible before 
any further steps are taken, because some reagents leave annoy- 
ing precipitates which must be removed, and others interfere 
with subsequent processes. Aqueous fixing agents with chromic 
acid as their principal ingredient are washed out with water ; 
aqueous solutions of corrosive sublimate are also washed out 
with water ; but alcoholic solutions should be washed out with 
alcohol of about the same strength as the fixing agent; picric 
acid, or fixing agents with picric acid as an ingredient, must not 
be washed out with water, but with alcohol, whether the picric 
acid be in aqueous or alcoholic solution. Running water is best, 
and where this is not convenient the water should at least be 
changed quite frequently. The washing-out process usually 
takes from twelve to twenty-four hours, but it can be shortened 
about one-half by keeping the fluid lukewarm. 

III. HARDENING AND DEHYDRATING. 

After the material has been washed, it is necessary to con- 
tinue the hardening and also to remove the water. Alcohol is 
used almost entirely for these purposes. It completes the 
hardening and at the same time dehydrates, that is, it replaces 
the water in the material, an extremely important considera- 
tion, for the least trace of moisture, a trace so slight as to be 
almost imaginary, is nevertheless sufficient to make a prepara- 
tion poor or indifferent when it might have been excellent. 

The process of hardening and dehydrating must be gradual. 
If the material should be transferred directly from water to 
absolute alcohol, the hardening and dehydrating would be 



General Method I 5 

brought about in a very short time, but the violent osmosis 
would cause a ruinous contraction of the more delicate parts. 
Therefore, transfer from water to 35 per cent, alcohol, which 
should act for six to twenty-four hours. Then use 50 per cent, 
for a similar period. Material may now be placed in 70 per 
cent, alcohol, where it may remain until ready for use, since 70 
per cent, alcohol is a good preservative. Various devices, like 
constant drips and osmotic apparatus, have been proposed to 
secure a more gradual transfer. Whether these have any real 
advantages still remains to be proved. The writer has taken 
well-fixed fern prothallia through the series 35 per cent., 50 per 
cent., 70 per cent., without the slightest plasmolysis. Such 
things as fern prothallia, filamentous algae, etc., can be watched 
under the microscope as the transfer is made, and, if plasmolysis 
results, the series of alcohols may be made closer, e. g. } 10 per 
cent., 20 per cent., 30 per cent., etc. It is said that material 
left for some time in 70 per cent, alcohol will shrink in spite of 
good killing and fixing, and it is also claimed that its capacity 
for staining is diminished. Some recommend that glycerine be 
added to the alcohol; others prefer to complete the dehydrating 
process and leave the material in an essential oil ; while still 
others would imbed it and keep it in paraffin. The last is doubt- 
less best of all, but requires such an immense amount of labor 
that it is impracticable for general purposes. Nearly all of our 
own material which is not needed for immediate use is in 70 
per cent, alcohol, unless, of course, the material has been put 
into formalin or some such reagent which kills, fixes, and pre- 
serves all at once. 

After the 70 per cent, alcohol, use 85 per cent, and 95 
per cent., allowing six to twenty-four hours for each. Then 
use 100 per cent, alcohol for one or two days. The 70 per 
cent, would probably complete all the hardening which is neces- 
sary, but the other three must be used to complete the removal 
of water. 

Up to this point the processes are exactly the same, whether 
the material is to be imbedded in parafrin or celloidin. 



1 6 Methods in Plant Histology 

IV. CLEARING. 

Let us suppose that the material has been thoroughly 
dehydrated, so that not the slightest trace of water remains. If 
the supposition chances to be contrary to fact, all the work 
which has preceded, as well as all which is to follow, is only an 
idle waste of time. The purpose of a clearing agent is to make 
the tissues transparent, but clearing agents also replace the 
alcohol. At this stage the latter process is the essential one, 
the clearing which accompanies it being incidental. The clear- 
ing, however, is very convenient, since it shows that the alcohol 
has been replaced and that the material is ready for the next step. 

Various clearing agents are in use. Xylol is the most gen- 
erally employed, and for most purposes it seems to be the 
best. Bergamot oil, cedar oil, clove oil, turpentine, and chloro- 
form are all necessary for special purposes. 

The transfer from absolute alcohol to the clearing agent 

should be gradual, like the hardening and dehydrating processes. 

The following is a good method : 

3 parts ioo per cent, alcohol and I part xylol, I to io hours. 
2 parts ioo per cent, alcohol and 2 parts xylol, I to io hours, 
i part ioo per cent, alcohol and 3 parts xylol, 1 to 10 hours. 

This transfer is best accomplished by adding the xylol to the 

alcohol. It is not necessary to measure, since anyone can guess 

with sufficient accuracy. Shake gently each time the xylol is 

added. Pour off the mixture and add pure xylol, which should 

cause the material to become transparent. This may require 

only a moment, but may require hours. Other clearing agents 

may be used in the same way instead of the xylol. 

V. THE TRANSFER FROM CLEARING AGENT TO PARAFFIN. 

This should also be a gradual process. The most con- 
venient method is to place a small block of paraffin in the pure 
clearing agent with the material. The paraffin dissolves gradu- 
ally and produces the same result as if a small shaving of 
paraffin had been added every few minutes for a day or so. 
During this process the bottle or dish should be kept lukewarm. 
Six to ten hours, or over night, is usually sufficient for this step, 



General Method ly 

although it would seem that material may be kept here for a 
much longer time without injury, and in case of such refractory 
material as the megaspores of the heterosporous pteridophytes 
four or five days seem necessary. Excellent preparations of the 
embryo-sac of Aster have been made from material which had 
remained in the xylol and paraffin for nearly three years. No 
more paraffin should be added than will go into perfect solution. 
The temperature may be gradually increased, so that a much 
greater amount of paraffin will go into solution, but the paraffin 
must not be allowed to crystallize. 

VI. THE PARAFFIN BATH. 

This step is usually called infiltration, but when the trans- 
fer from the clearing fluid to paraffin is made gradually, as has 
just been indicated, the process of [infiltration is already begun. 
It is nowfnecessary to get rid of the xylol or other clearing 
agent. This may be done by simply pouring off the mixture of 
xylol iand paraffin and replacing it with pure melted paraffin. 
The bath should be kept at a temperature about i° C. higher 
than the melting-point of the paraffin. Fifty-three degrees C. 
is a good temperature for general purposes, but this may be 
reduced from i° to 3 C. in winter, and must often be raised 
in summer. For special purposes it is sometimes necessary to 
use a temperature as high as jo° C. If the xylol or other 
clearing agent is not thoroughly removed, the paraffin will be 
granular or mealy, and will not cut well. The paraffin should be 
changed once or twice to make sure that the clearing agent 
is all removed. Do not waste this paraffin, for the clearing 
agent can be driven off by prolonged heating, and the paraffin 
will be better than ever. Most people use soft paraffin (about 
45 C.) for the first half of the time necessary for infiltration, 
and a harder grade (48 C. to 54 C.) for the latter part of the 
process. This is a good plan, for soft paraffin melts at a lower 
temperature, and it is always well to minimize heat. 

Some think that it is better not to pour off the mixture of the 
clearing agent and paraffin, but rather to evaporate the clearing 



1 8 Methods in Plant Histology 

agent by keeping the temperature just high enough to prevent 
the crystallization of the paraffin. This method has certainly 
given good results with small, delicate objects. 

The time required for infiltration varies with the character 
of the tissue and the size of the piece. Few things can be well 
infiltrated in less than an hour. Lily ovaries require one to four 
hours ; heads of Aster at the fertilization period, six to twelve 
hours. Some claim that even delicate objects, like fern pro- 
thallia and the filamentous algae and fungi, are not injured by 
a bath of several days, if care be taken not to let the tempera- 
ture rise above 48° C. to 50 C. No one seems to know how 
long a certain object should remain in the bath, but many 
competent investigators are now using more prolonged periods. 

VII. IMBEDDING. 

Material may be imbedded in paper trays, watch crystals, 
or in apparatus made for the purpose. Imbedding |_s consist- 
ing of two L-shaped pieces {fig. 11) of brass, 
type metal, or lead are very convenient. 
We use a pair-of L-shaped pieces with arms 
three inches long. These furnish a box of 
almost any required size. A piece of glass 
serves for a bottom. The tray, minot, or 
whatever is used, should be slightly smeared 
with glycerine, to prevent sticking. If several objects are to be 
imbedded in one dish, it is best to have the dish as near the 
temperature of melted paraffin as possible ; otherwise the objects 
may stick to the bottom, and it will be impossible to arrange 
them properly. Great care should be taken, however, not to 
have the dish too hot, since too high a temperature not only 
injures the material, but also prevents a thorough imbedding. 
Pour the paraffin with the objects into the imbedding dish and 
cool as rapidly as possible. If paraffin cools slowly, it crystal- 
lizes and does not cut well. The layer of paraffin should be 
just thick enough to cover the objects, not only as a matter of 
economy, but because a thick layer retards the cooling. Very 




General Method 



19 



small objects, like the megaspores of Marsilea, ovules of Silphium, 
etc., may simply be poured out upon a cool piece of glass. In 
this way very thin cakes are made, which harden very rapidly. 

VIII. CUTTING. 

Twenty minutes after an object is imbedded it is ready for 
cutting. Trim the paraffin containing the object into a conveni- 
ent shape, and fasten it upon a block of wood. Blocks of pine 
three-fourths of an inch long and three-eighths of an inch square 
are good for general purposes. Put paraffin on the end of the 
block so as to form a firm cap about one-eighth of an inch thick. 
Warm the cap and the bottom of the piece containing the object, 
and press them lightly together ; then touch the joint with a hot 
needle, put the whole thing into cold water for a minute, and it 
is ready for cutting. Cutting can be learned only by experience, 
but a few hints may not come amiss : [a) Keep the knife sharp. 
If expense is not too serious an objection, it is well to have two 
hones, one rather soft, for use when the knife is dull, and the 
other quite hard, for putting on an even edge. Flood the stone 
with water, and rub it with the small slip which accompanies all 
high-grade hones ; this not only makes a lather which facilitates 
the sharpening, but it also keeps the surface of the hone flat. 
As soon as the edge of the knife appears smooth and even 
under a magnification of thirty or forty diameters, the sharpen- 
ing is completed with a good strop. It is better to sharpen the 
knife every time you use it. {b) Keep the microtome well oiled 
and clean, (c) Trim the block so that each section shall be a 
perfect rectangle. 



B 
Fig. 12. 



20 Methods in Plant Histology 

A ribbon of sections like that shown in fig. 12 A is much 
better than one like B of the same figure, because sections will 
usually come off in neater ribbons if the knife strikes the longer 
edge of the rectangle, so that the sections are united by their 
longer sides rather than by the shorter. Crooked ribbons are 
caused by wedge-shaped sections, and are always to be avoided, 
because they make it difficult to economize space, and also 
because they present such a disorderly appearance. The knife, 
which should be placed at a right angle to the block and not 
obliquely, should strike the whole edge of the block at once, and 
should leave in the same manner. 

IX. FIXING SECTIONS TO THE SLIDE. 

Sections must be firmly fixed to the slide, or they will be 

washed off during the processes involved in staining. Mayer's 

albumen fixative is excellent for this purpose. Formula : 

White of egg, 50 cc. (Active principle.) 

Glycerine, 50 cc. (To keep it from drying up.) 

Salycilate of soda, 1 g. (Antiseptic, to keep out bacteria, etc.) 

Shake well and filter. It will keep from two to six months. 

The fixative may be used alone or in connection with the water 

method. Put a small drop of fixative on the slide, smear it 

evenly over the surface, and then wipe it off with a clean finger 

or piece of linen until only a scarcely perceptible film remains ; 

then add several drops of distilled water and float the sections 

or ribbons on the water. Warm gently until the paraffin 

becomes smooth and free from wrinkles. Be careful not to 

melt the paraffin, for the albumen of the fixative coagulates 

with less heat than is required to melt the paraffin. It is a very 

good plan to put the slide on the top of the water bath for a 

moment and then, after the sections have become smooth, 

remove the surplus water and leave them on the bath with a 

couple of thicknesses of blotting paper under them for three or 

four hours, or, better, over night. If the fixative is used alone, 

as is often the case when sections are very thick, none of this 

delay is necessary, since the sections are merely laid upon the 

fixative and pressed down gently with the finger. 



General Method 21 

X. REMOVAL OF THE PARAFFIN. 

To remove the paraffin it is very customary to heat the 
slide gently until the paraffin melts, and then place the slide in 
xylol for thirty seconds or a minute. I believe, however, that 
it is far better merely to warm the slide a little (not warmer than 
30° C). Good xylol will then remove the paraffin in two or 
three minutes. Even if the slide should not be warmed at all, 
good xylol should remove the paraffin in five minutes. 

XI. REMOVAL OF XYLOL. 

The xylol may be removed either by absolute alcohol 
or by the 95 per cent. The absolute alcohol does not seem to 
be really necessary. For my own work I have two stender 
dishes of xylol and two of absolute alcohol. After the xylol 
has been used for a time I employ it only for removing paraffin, 
and in the same way use the absolute alcohol only for removing 
the xylol, while the other two dishes are used only for dehydrat- 
ing and clearing. In transferring from xylol to absolute alcohol, 
or vice versa, it is well to drain off the superfluous liquid by 
resting the corner of the slide upon a piece of blotting paper. 
These rather expensive reagents will last much longer when this 
precaution is taken. 

XII. TRANSFER TO THE STAIN. 

Stains are aqueous or alcoholic, and alcoholic stains are 
of various strengths. If an aqueous stain be used, the slide 
should be passed successively through the alcohols 95 per cent., 
85 per cent., 70 per cent., 50 per cent., and 35 per cent., allow- 
ing each to act for about thirty seconds, after which the slide is 
put into the stain. (In many cases it is sufficient to put a few 
drops of the stain on the slide with a pipette.) From the stain 
the slide is passed back through the various grades of alcohol, 
allowing it to remain about thirty seconds in each as before. If 
the stain be alcoholic of about 70 per cent, strength, the process 
is somewhat shorter, for the slide goes into the stain from 70 
per cent, alcohol, and goes back into 70 per cent, alcohol from 
the stain. The rule is to transfer to the stain from the alcohol 



22 Methods in Plant Histology 

which is nearest the strength of the stain. These directions are 
based upon a rather wide experience and certainly give excellent 
results with stains which do not forbid their application. How- 
ever, evidence has long been accumulating which indicates that 
in case of very thin sections (sections so thin that no whole cells 
are included) only the absolute alcohol is necessary and that 
the rest may be omitted. Whether it is better to omit the 
alcohols from 35 per cent, to 95 per cent, when staining on the 
slide is a question still waiting for a definite answer, although 
careful and systematic experimenting would soon settle it. 
Even if it should be shown that it is as well or better to omit 
the alcohols from 35 per cent, to 95 per cent, in case of thin 
sections, it must not be inferred that these alcohols may be 
omitted in preparing a piece of epidermis stripped from a leaf, 
for, in this case, a ruinous plasmolysis occurs, and in sections 
thick enough to include whole cells the same damage is done. 
In the directions given in this book it is assumed that the series 
35 per cent., 50 per cent., 70 per cent., 85 per cent., 95 per cent., 
and IOO per cent, is to be used, unless otherwise mentioned. 

XIII. CLEARING. 

After the sections have been stained, passed back through 
the various grades of alcohol, and have been thoroughly dehy- 
drated in absolute alcohol, they are cleared or made transparent 
by means of xylol or some other clearing agent. The clearing 
agent must be a solvent of balsam. From thirty seconds to five 
minutes will be sufficient for clearing any kind of sections. 

XIV. MOUNTING IN BALSAM. 

After the sections are cleared, wipe the slide on the side 
which does not bear the sections. Put on a drop of Canada 
balsam and add a clean, 1 thin cover. Before the cover is put 
on, pass it through the flame of an alcohol lamp to remove 
moisture, for it would be a pity indeed to injure a preparation 

1 Slides and covers should be treated with hydrochloric acid, or equal parts of 
hydrochloric acid and water, for several hours. They should then be thoroughly rinsed 
in water and placed in 95 per cent, alcohol. They should be wiped with a cloth per- 
fectly free from lint. 



General Method 23 

at this stage of the process. Add a label, and the mount is 
complete. 

A TENTATIVE SCHEDULE FOR PARAFFIN SECTIONS. 

It will be useful to give several tentative schedules for the 
use of beginners. It cannot be too strenuously insisted that 
these schedules are only tentative, their sole object being to give the 
beginner a start. The following is a tentative schedule for the 
ovary of a lily at any period before fertilization. The pieces 
should not be more than half an inch in length. 

1. Chromo-acetic acid, 2 days. 

2. Wash in water, 1 day. 

3. Thirty-five per cent., 50 per cent., 70 per cent., 85 per cent., 95 per cent, 
alcohol, 6 to 24 hours each, as convenient. 

4. One hundred per cent, alcohol, 24 hours. This should be changed once 
or twice. The volume should be at least ten times that of the material. 

5. Transfer from absolute alcohol to xylol, allowing at least 2 hours in each 
of the three mixtures, and 2 hours in pure xylol. 

6. Add paraffin to the xylol and keep warm for 12 to 24 hours. 

7. Melted paraffin in the bath, 2 to 24 hours, as convenient. The paraffin 
should be changed once or twice. 

8. Imbed. 

9. Section; about 10 /x is a good thickness. 

10. Fasten to the slide. 

11. Dissolve off the paraffin in xylol. 

12. One hundred per cent., 95 per cent., 85 per cent., 70 per cent., 50 per 
cent, alcohol, 30 seconds each. An hour in each would do no damage 
if other duties should interfere. 

13. Delafield's hematoxylin, 10 minutes. 

14. Rinse in water, 5 minutes. A couple of hours does no harm. 

15. Thirty-five per cent., 50 per cent., and 70 per cent, alcohol, 30 seconds 
each. An hour would do no harm. 

16. Acid alcohol, 1 second. 

17. Seventy per cent, alcohol, 1 minute. An hour would do no harm. 

18. Erythrosin, 30 seconds to 1 minute. 

19. Eighty-five per cent., 95 per cent., 100 per cent., about 5 seconds each. 
This step must not be too prolonged, lest the erythrosin wash out. 

20. Xylol, at least 1 minute. The slide may be left here for an hour without 
injury. 

21. Mount in balsam. 



24 Methods in Plant Histology 

On the whole, it is not a good plan to use protractedjperiods 
in the processes from No. 1 1 to No. 20 inclusive, because there 
is danger that a prolonged soaking may loosen some of the more 
delicate contents of the cells, even if whole sections do not 
come off bodily. 



CHAPTER V. 

KILLING AND FIXING AGENTS. 

In this short account only the reagents which are at present 
considered most valuable for botanical work will be considered. 
Probably no process in microtechnique is in more urgent need 
of improvement than this first step of killing and fixing. Nearly 
all of our formulae are merely empirical, for very few botanists 
are expert chemists, and those who have the requisite knowl- 
edge of chemistry are interested in physiological problems 
rather than in microtechnique. The principal ingredients of the 
usual killing and fixing agents are : alcohol, chloroform, chromic 
acid, acetic acid, osmic acid, formic acid, picric acid, sulphuric 
acid, platinum chloride, iridium chloride, corrosive sublimate, 
and formalin. We shall consider first 

THE ALCOHOLS. 

a. Ninety-five per cent. Alcohol. — This is in quite general use 
for material which is needed only for rough work. It is extremely 
convenient, since it kills, fixes, and preserves at the same time 
and needs no changing or washing. It really has nothing to 
recommend it for fine work. It causes protoplasm to shrink, but 
cell walls usually retain their position, so that 95 per cent, alco- 
hol will do for free-hand sections of wood and many herbaceous 
stems ; but even free-hand sections of tender stems, like young 
geraniums and begonias, will look better if better reagents are 
employed. Alcohols weaker than 95 per cent, are not to be 
recommended as fixing agents, although 70 per cent, alcohol, or 
even 50 per cent., will preserve material for habit work. 

b. Absolute (100 per cent.) Alcohol. — This is a fair killing and 
fixing agent, but is rather expensive. It causes but little shrink- 
ing of the protoplasm, and is a time-saver if material is to be 
imbedded in paraffin. With 95 per cent, or with absolute 

25 



26 Methods in Plant Histology 

alcohol, objects are generally left in the reagent until needed for 
use, but such material becomes very brittle. The addition of 
glycerine is an improvement, if material is to be kept long. 
Acetic acid has been used with alcohols to counteract the 
tendency to shrink. One of the most successful of the alcohol 
combinations is 

c. Carnoy's Fluid. — 

Absolute alcohol, 6 parts. 

Chloroform, 3 parts. 

Glacial acetic acid, 1 part. 

The penetration of the reagent is excellent, and only a few 
hours are needed for fixing. Material should be washed in 
absolute alcohol (perhaps 95 per cent, alcohol would do no 
harm) until there is no odor of acetic acid. This should not 
require more than one or two hours. It is better to imbed in 
paraffin at once, but when this is not convenient the material 
may be transferred to 85 per cent, alcohol and then to 70 per 
cent., where it maybe left until needed. Cyanin and erythrosin, 
fuchsin and iodine green, and similar combinations, give particu- 
larly brilliant staining after this reagent. 

THE CHROMIC-ACID GROUP. 

Chromic acid, or solutions with chromic acid as a foundation, 
are the most generally useful killing and fixing agents yet known 
to the botanist. A 1 per cent, solution of chromic acid in water 
gives good results, but it is better to use the chromic in connec- 
tion with other ingredients, such as acetic acid, formic acid, 
osmic acid, etc. The proportions of the various ingredients, 
for the present at least, must be determined by experiment. 
With favorable objects like fern prothallia, Spirogyra, and other 
things which can be watched while the fixing is taking place, 
suitable proportions are rather easily determined, because speci- 
mens, after being placed in the reagent, may be examined at 
frequent intervals, and combinations which cause plasmolysis 
may be rejected and different proportions tried until satisfactory 
results are secured. For example, fern prothallia might be placed 
in the following solution : chromic acid, 2 g.; acetic acid, 1 cc; 



Killing and Fixing Agents 27 

and water, 97 cc. If plasmolysis takes place, weaken the chromic 
or strengthen the acetic, since the chromic has a tendency to 
produce contraction, and the acetic to cause swelling. Too 
large a proportion of acetic acid, however, may cause distortion, 
and hence it would be better to weaken the chromic. In case 
of fern prothallia, 3 parts chromic, 1 part acetic, and 396 parts 
water will cause practically no plasmolysis, and the fixing is 
sufficiently thorough to permit imbedding in paraffin. A com- 
bination may be quite satisfactory for fern prothallia and still 
fail to give good results with Spirogyra, and a combination which 
succeeds very well with Spirogyra may not succeed at all with Van- 
cheria. For very critical work the most favorable proportions 
must be determined for the particular plant under investigation. 
When the effect of the reagent cannot be observed directly, it is 
well to make a free-hand section and thus determine whether 
plasmolysis takes place. It is not safe to judge the action of a 
fixing agent by the appearance of sections cut from material 
which has been imbedded in paraffin, because shrinking of the 
cell contents often takes place during the transfer from absolute 
alcohol to the clearing agent or during infiltration with paraffin, 
and sometimes during even later processes. When in doubt as 
to proportions, we should suggest 2 cc. chromic acid, 2 cc. acetic 
acid, and 296 cc. water as a good formula for most purposes. 

The time required for fixing undoubtedly varies with different 
plants, but twenty-four hours may be considered a minimum 
even for the most delicate objects. It is well known that zoolo- 
gists allow fixing agents like Miiller's fluid and Erlicki's fluid to 
act for weeks before the material is passed on to the next 
stage, and it may well be questioned whether botanists have 
not made a mistake in allowing the chromic solutions to act for 
so short a time. At present most botanists recommend sixteen 
to twenty-four hours for material which is to be imbedded in 
paraffin, but some recent experiments in my laboratory indicate 
that material which has been in the fixing fluid for two or three 
days is better able to withstand the subsequent processes. More 
rapid penetration, and consequently more immediate killing, can 



28 Methods in Plant Histology 

be secured if the reagent is kept warm (30 to 40 C). Since 
chromic acid has a tendency to render objects hard and brittle, 
it is often better to use some other fixing agent, if much diffi- 
culty is anticipated in the cutting. 

After fixing is complete, the reagent should be washed out 
with water. Running water is desirable, and where this is not 
convenient the water must be changed frequently. Any material 
should be sufficiently washed in six to twenty-four hours, but 
the time may be shortened about one-half by using lukewarm 
water. 

Some of the formulae are as follows : 

a. Strong Chromo-Acetic Solution. — 

1 g. chromic acid. 

1 cc. glacial acetic acid. 

98 cc. water. 

This solution has been used quite extensively in embryo- 
logical work upon the higher plants. 

b. Weak Chromo-Acetic Solution. — (Schaffner's formula): 

0.3 g. chromic acid. 
0.7 acetic acid. 

99 cc. water. 

This has also been used in embryological work. It causes 
little or no plasmolysis, but the chromic seems rather weak. 
Difficult material, like Aster heads and ripe Capsella pods, cuts 
more readily after this reagent than after the stronger solution. 

c. Medium Chromo-Acetic Solution. — 

0.7 g. chromic acid. 

0.5 cc. glacial acetic acid. 

100 cc. water. 

.-> 

We are now using this solution, and it seems very ^successful 
in nearly all cases. 

d. Flemming's Fluid. — (Weaker solution.) 

( 1 per cent, chromic acid, 25 cc. 

A. < 1 per cent, acetic acid, 10 cc. 
I Water, 55 cc. 

B. 1 per cent, osmic acid, 10 cc. 



Killing and Fixing Agents 29 

Keep the mixture A made up, and add B as the reagent is 
needed for use, since it does not keep well. This fluid is quite 
expensive on account of the osmic acid. For cytological work 
it gives as good results as any fixing agent which has yet been 
thoroughly tested. It is especially recommended for chromo- 
somes, centrosomes, achromatic structures, and mitotic phe- 
nomena in general. Material should be in very small pieces 
one-eighth of an inch square, or in thin slices one-eighth of an 
inch or less in thickness, for the fluid penetrates poorly. The 
blackening due to the osmic acid may be removed by peroxide 
of hydrogen just before the slide is passed from the alcohol into 
the stain. Flemming's safranin-gentian violet-orange combina- 
tion gives its most brilliant results after this reagent. It seems 
possible that the traditional superiority of this reagent has been 
overestimated. 

e. MerkePs Fluid. — 

Equal volumes of a 1.4 per cent, solution of chromic acid and a 1.4 per 
cent, solution of platinic chloride. 

This is also an expensive reagent. It is recommended for 
mitotic phenomena, but does not seem to equal Flemming's 
solution. 

/ Hermann's Fluid. — 

1 per cent, platinic chloride, 15 parts. 
Glacial acetic acid, 1 part. 

2 per cent, osmic acid, 4 or 2 parts. 

This is the most expensive fixing agent yet discovered, and 
for botanical purposes it does not seem to be any better than the 
cheaper chromic mixtures. It is mentioned here with chromic 
mixtures because it originated as a variation of Flemming's 
fluid, the platinic chloride being substituted for the chromic 
acid. 

According to Lee, the chief objection to all mixtures into 
which chromic acid enters is that "it precipitates certain of the 
liquid albuminoids of the tissues in the form of filaments or net- 
work, which are often of great regularity and simulate structural 
elements of the tissues." Nevertheless, the mixtures which have 



30 Methods in Plant Histology 

just been described are the best which have yet been thoroughly 
tested. If material killed in any of the above mixtures is not 
well washed, the hematoxylins will not stain. It is claimed that 
the anilins will stain in spite of poor washing, but it is a question 
whether such preparations are as permanent as those from well- 
washed material. 

PICRIC ACID. 

Use a saturated solution in water or 70 per cent, alcohol. 
One gram of picric acid crystals will saturate about 75 cc. of 
water or alcohol. This reagent penetrates well and does not 
make the material brittle. It is to be recommended when diffi- 
culty is anticipated in the cutting. If used cold, the time varies 
from one to twenty-four hours, depending upon the character of 
the tissue and size of the specimen. If used hot (85 C), five 
or ten minutes will be sufficient. Material should be washed in 
70 or 50 per cent, alcohol. Water is injurious, and some even 
go so far as to avoid aqueous stains, unless the material has been 
thoroughly washed. The washing should be continued until the 
material appears whitish and the alcohol no longer becomes 
tinged with yellow. Picro-carmine gives its best results after 
this reagent. Picric acid can be combined with various other 
fixing agents, and so we have picro-sulphuric acid, picro-nitric 
acid, picro-chromic acid, picro-chromic-sulphuric acid, and picro- 

osmic acid. 

CORROSIVE SUBLIMATE. 

Use a 2 to 5 per cent, solution in water, or 70 per cent, alco- 
hol. The addition of about 1 cc. of glacial acetic acid to 100 cc. 
of this solution is certainly an improvement. The time required 
is considerably shorter than for chromic solutions. From one to 
ten hours will be found to be sufficient. If used hot (85 ° C), 
only five or ten minutes is required. Washing may be done 
with water, but 50 per cent, alcohol is better. If a few drops of 
an iodine solution be added to the alcohol, the alcohol takes on 
a brownish color, but soon clears up. If the addition of iodine 
be continued, the washing is complete when the alcohol no 
longer clears up, but retains the brown color. If the washing is 



Killing- and Fixing Agents 31 

incomplete, crystals of corrosive sublimate will be unduly con- 
spicuous in preparations made from the material. Camphor may 
be used instead of iodine to hasten the washing. 

The carmines are very brilliant after corrosive sublimate on 
account of the formation of mercuric carminate, but haematoxy- 
lin and anilines also give good results. It is claimed, however, 
that achromatic structures do not stain well with the safranin-gen- 
tian violet-orange combination after this fixing agent, but it 
might be worth while to test other stains before discarding this 
reagent for cytological work. 

Corrosive sublimate material gets very brittle if allowed to 

remain long in alcohol, and therefore it is better to imbed it as 

soon as possible. 

FORMALIN. 

Formalin is a comparatively recent addition to the list of 
killing and fixing agents. It is an excellent preservative, often 
preserving the blue and red colors as well as the structure of 
objects. A 2 or 4 per cent, solution in water is good for fila- 
mentous algae. The material may simply be put into the reagent 
and left until needed for use. For class use formalin material 
should be washed in water for several minutes, because the 
fumes are irritating to the eyes and mucous membranes. After 
a thorough washing in water any of the usual stains may be 
used. 

GENERAL HINTS ON FIXING. 

It is very desirable that the fixing agent should penetrate 
quickly to all parts of the object. For this reason material 
should be in small pieces. The best fixing agents do their best 
work near the surface of the piece. Small objects like Azolla 
and fern prothallia may be thrown into the fixing agent entire ; 
even larger objects, which, like the anthers of Liliurn, are easily 
penetrated, may also be put in without any cutting. Most 
objects larger than quarter of an inch cubes should be trimmed 
with a sharp knife or razor ; some knowledge of the structures 
concerned is essential before one can trim material with unvary- 
ing success. 



32 Methods in Plant Histology 

Some objects, although small, cause trouble in various ways. 
Many buds are hairy and will not sink ; if such things are dipped 
quickly in strong alcohol, they will usually sink. If rather large 
air bubbles prevent the material from sinking, as in case of peri- 
chaetical leaves of some mosses and involucral leaves of liver- 
worts, a little dissection or a careful snip with the scissors will 
obviate the difficulty. If an air-pump is available, the bubbles 
are easily removed. 

It is often asked whether fixing agents really preserve the 
actual structure of cell contents. It must be admitted that 
some things, notably the liquid albuminoids, are much modified 
in appearance, but the most competent observers are now 
inclined to believe that such delicate objects as chromosomes, 
centrosomes, the achromatic figure, and even the structure of 
protoplasm, can be studied with confidence from material which 
has been fixed, imbedded, and stained. Recent study of these 
objects in the living condition has strengthened this confidence. 

It is certain that we have not yet found the ideal fixing 
agent for cell contents. Such an agent must not be a solvent 
of any of the cell contents, must penetrate rapidly, must pre- 
serve structures perfectly, and must harden so thoroughly that 
every detail shall remain unchanged during the subsequent 
processes of dehydrating, clearing, imbedding, sectioning, and 
staining. 



CHAPTER VI. 

STAINING. 

Staining is one of the most important and most complicated 
processes of microtechnique. The formulae are largely empirical, 
and there is still abundant room for experimentation in the use 
of mordants and in the effect of the same stain or combination 
after various fixing agents. 

Stains may be classified in various ways ; e.g., there are three 
great groups of stains — the Carmines, the Hematoxylins, and 
the Anilins. Stains may be classified as basic and acid, or they 
may be regarded as general and specific. A general stain affects 
all the elements, while a specific stain affects only certain 
elements or stains some elements more deeply than others. 
Stains which show a vigorous affinity for the nucleus have been 
called nuclear stains, and those which affect the cytoplasm more 
than the nucleus have been termed plasma stains. Of course, 
such stains are specific. 

We shall consider some of the more important hematoxylins, 
carmines, and anilins, reserving general directions and theoreti- 
cal questions until the end of this chapter. Many of the formulae 
are taken from The Microtomisfs Vade-Mecum (Lee), which is 
easily the most complete compendium of stains and other 
reagents concerned in microtechnique. It is to be regretted 
that botanists have no book of this character, but it must be 
confessed that we have not the material for such an extensive 
work. Other formulae are from Botanical Microtecluiiqne (Zim- 
mermann) and from Stirling's Histology. The directions for 
using a stain apply to stains made up according to the formulae 
which are given here, and may need modification if other formulae 
are employed. It is hoped, however, that the directions will 
give the student sufficient insight into the rationale of staining 
to enable him to make any necessary modifications. 

33 



34 Methods in Plant Histology 

THE HEMATOXYLINS. 

The most important hematoxylins are Delafield's hema- 
toxylin, Kleinenberg's hematoxylin, Erlich's hematoxylin, 
Boehmer's hematoxylin, Mayer's hem-alum, and Haidenhain's 
iron alum-hematoxylin. 

All the hematoxylins mentioned above contain alum, and, 
according to Mayer, who has written the most important work 
on hematoxylin stains ("Ueber das Farben mit Hematoxylin," 
Mitth. a. d. Zool. Station zu Neapel, 10 : 170-186, 1891, and "Ueber 
Hematoxylin, Carmin und verwandten Materien," Zeit. f. wiss. 
Mikr., 16: 196-220, 1899). "The active agent in them is a com- 
pound of hematin with alumina. This salt is precipitated in the 
tissues, chiefly in the nuclei, by organic and inorganic salts there 
present (e. g., by the phosphates) and perhaps also by other 
organic bodies belonging to the tissues." These salts are fixed 
in the tissues by the killing and fixing agent, and when the stain 
is applied a chemical combination results. Hematoxylins stain 
well after any of the fixing agents described in the preceding 
paper, but they are most effective when used after members of 
the chromic-acid series. 

Delafield's Haematoxylin. — "To 100 cc. of a saturated solution 
of ammonia alum add, drop by drop, a solution of I g. of hema- 
toxylin dissolved in 6 cc. of absolute alcohol. Expose to air 
and light for one week. Filter. Add 25 cc. of glycerine and 
25 cc. of methyl alcohol. Allow to stand until the color is 
sufficiently dark. Filter, and keep in a tightly stoppered bottle." 
(Stirling and Lee.) The addition of the glycerin and methyl 
alcohol will precipitate some of the ammonia alum in the form 
of small crystals. The last filtering should take place four 
or five hours after the addition of the glycerine and methyl 
alcohol. 

The solution should stand for at least two months before it 
is ready for using. This "ripening" is brought about by the 
oxidation of hematoxylin into hematin, a reaction which may 
be secured in a few minutes by a judicious application of per- 
oxide of hydrogen. 



Stai?iing 3 5 

Transfer to the stain from 35 per cent, alcohol or from water. 
The length of time required is exceedingly variable. Sometimes 
sections will stain deeply in three minutes, but it is often necessary 
to stain for thirty minutes or even longer. This stain may be 
diluted with several times its own volume of water ; when this is 
done, the time required is correspondingly long, but the staining 
is frequently more precise. The length of time required will be 
fairly uniform for all material taken from the same bottle. This 
fact indicates that the washing process, which follows killing and 
fixing, is an important factor; if the washing has been thorough, 
the material will stain readily; but if the washing has been insuf- 
ficient, the material may stain slowly or not at all. The washing 
is particularly important when the fixing agent contains an acid. 
Transfer from the stain to water. Distilled water is neither 
necessary nor desirable. Some writers recommend washing for 
twenty-four hours, but this seems unnecessary; an hour is usually 
enough, and a few minutes is often sufficient. Precipitates are 
often formed when slides are transferred directly to alcohol from 
this stain ; otherwise, it would be better to transfer from the 
stain to 35 per cent, alcohol. Pass through the alcohols to 70 
per cent, alcohol and then give the slide a few dips (two seconds 
is often sufficient) in acid alcohol (0.5 cc. HC1. to 100 cc. of 
70 per cent, alcohol). This extracts the stain more rapidly from 
other parts than from the nuclei, and hence gives a good nuclear 
stain. Some prefer to stain for a very short time and use no 
acid alcohol, but, as a rule, it seems best to overstain and then 
differentiate in this way, because sharper contrasts are obtained. 
Transfer from acid alcohol to 70 per cent, alcohol and leave here 
until a rich purple color replaces the red due to the acid. Since 
small quantities of the acid alcohol are carried over into the 70 
per cent, alcohol, it is well to add a drop of ammonia now and 
then to neutralize the effect of the acid. Too much ammonia is 
to be avoided, for it gives a disagreeable bluish color with poor 
differentiation, probably on account of the precipitation of 
alumina. The slide may now be passed through the alcohols, 
cleared in xylol, and mounted in balsam ; or, if a double stain be 



36 Methods in Plant Histology 

preferred, treat for thirty seconds to one minute with eosin, 
erythrosin, or some other stain affording a good contrast; rinse 
in 70 per cent, alcohol and proceed as usual. 

Delafield's haematoxylin is the most generally useful stain in 
the haematoxylin group. It brings out cellulose walls very 
sharply, and consequently is a good stain for embryos and the 
fundamental tissue system in general. With safranin it forms a 
good combination for the vascular system, the safranin giving 
the lignified elements a bright red color, while the haematoxylin 
stains the cellulose a rich purple. It is a good stain for chro- 
matin, and the achromatic structures show up fairly well, but can 
be brought out much better by special methods. Archesporial 
cells and sporogenous tissue are very well defined if proper care 
be taken. Whenever you are in doubt as to the selection of a 
stain for general purposes, we should advise the use of Delafield's 
haematoxylin. 

The following is a general schedule for staining paraffin sec- 
tions on the slide in Delafield's haematoxylin : 

1. Stain (from water or 35 per cent, alcohol) 5 minutes. 

2. Rinse in water, 5 minutes. 

3. Thirty-five per cent., 50 per cent., and 70 per cent, alcohol, 30 seconds 
each. 

4. Acid alcohol, 1 second. 

5. Seventy per cent, alcohol, 1 minute. 

6. Eighty-five per cent., 95 per cent., and 100 per cent, alcohol, 30 seconds 
each. 

7. Xylol, 1 minute. 

8. Mount in balsam. 

If, after rinsing in water, the stain is evidently too weak, put 
the slide or section back into the stain until it appears over- 
stained. For thin sections (lOAt or less) thirty seconds in each 
of the alcohols will be unnecessarily long, although thirty minutes 
would do no damage. Give the slide a single dip into the acid 
alcohol, transfer it quickly to the 70 per cent, alcohol, and then 
examine it ; if it still appears overstained, give it another dip in 
the acid alcohol, and repeat the process until the stain is what 
you want. After the haematoxylin is just right, apply a contrast 



Staining 3 7 

stain, if you wish to double-stain. It is a good plan to move the 
slide gently to and fro in the absolute alcohol. Before trans- 
ferring to the xylol, wipe the alcohol from the back of the slide, 
or at least rest the corner of the slide upon blotting paper 
for two or three seconds, in order that you may not carry over 
so much alcohol into the xylol, and thus impair this rather 
expensive reagent. The slide may also be moved gently to and 
fro in the xylol. Add a drop of balsam and a cover. Since the 
xylol is very volatile, this last step must be taken quickly. If 
blackish spots appear, they are usually caused by the drying of 
sections before the balsam and cover are added ; if there are 
whitish spots or an emulsion-like appearance, the clearing is not 
thorough; this may be caused by poor xylol (or other clearing 
agent); by absolute alcohol which is considerably weaker than 
its name implies (the absolute alcohol must test at least as high 
as 99 per cent., and ought to test as high as 99.5 per cent., if 
xylol is to be used for clearing) ; or the difficulty may be caused 
by passing too quickly through the absolute alcohol and xylol, 
or may even be caused by moisture on the cover-glass. 

Kleinenberg's Hematoxylin.— This stain has had a wide use, 
but it is now largely replaced by better formulae. It is men- 
tioned here merely because it is on the shelves of so many 
laboratories. It is doubtful whether it is equal to Delafield's in 
any kind of botanical work. A good description is given in the 
Quarterly Journal of the Microscopical Society, 74 : 208 — , 1897. 

Erlich's Hematoxylin. — 

Distilled water, 50 cc. 
Absolute alcohol, 50 cc. 
Glycerine, 50 cc. 
Glacial acetic acid, 5 cc. 
Hematoxylin, 1 g. 
Alum in excess. 

Keep it in a dark place until the color becomes a deep red. 
If well stoppered, it will keep indefinitely. Transfer to the stain 
from 50 per cent, or 35 per cent, alcohol. Stain five to thirty 
minutes. Since there is no danger from precipitates and the 



(Al 



38 Methods in Plant Histology 

solution does not overstain, it is not necessary to treat with 
water or with acid alcohol, but the slide may be transferred from 
the stain to 70 per cent, alcohol. Eosin, erythrosin, or orange 
G are good contrast stains. 
Boehmer's Hematoxylin, — 

j Hematoxylin, 1 g. 
Absolute alcohol, 12 cc. 
Alum, 1 g. 
•istilled water, 240 cc. 

The solution A must ripen for two months. When wanted 
for use, add about 10 drops of A to 10 cc. of B. Stain ten to 
twenty minutes. Wash in water and proceed as usual. 

Mayer's Haem-Alum. — Hematoxylin, 1 g., dissolved with 
heat in 50 cc. of 95 per cent, alcohol and added to a solution 
of 50 g. of alum in a liter of distilled water. Allow the mixture 
to cool and settle ; filter ; add a crystal of thymol to preserve 
from mold. (Lee.) 

It is ready for use as soon as made up. Unless attacked by 
mold, it keeps indefinitely. Transfer to the stain from water. 
It is seldom necessary to stain for more than ten minutes, and 
four or five minutes is generally long enough. As a rule, better 
results are secured by diluting the stain (about 1 cc. to 10 cc. of 
distilled water) and allowing it to act for ten hours or over night. 
This is a good stain for the nuclei of filamentous algae and fungi, 
since it has little or no effect upon cell walls or plastids. Wash 
thoroughly in water and transfer to 10 per cent, glycerine. 
Specimens may be mounted in balsam, if they can be gotten 
through without shrinking. 

Haidenhain's Iron Alum-Haematoxylin. — This stain was intro- 
duced by Haidenhain in 1892 and has gained a well-deserved 
popularity with those engaged in cytological work. Two solu- 
tions are used, and they are never mixed : 

A. One and a half to 4 per cent, aqueous solution of ammonia sulphate 
of iron. (At present we use a 3 per cent, solution.) 

B. One-half per cent, aqueous solution of hematoxylin. 

The first solution acts as a mordant, i. e., it does not stain, 
but prepares the tissue for the action of the second solution. 



Staining 39 

Transfer to the iron-alum from water ; allow this solution to 
act for two hours ; wash in water five minutes, and then stain 
in the y 2 per cent, hematoxylin ten hours or over night. 
Rinse in water five minutes and treat for a second time with the 
iron alum, which now rapidly extracts the stain. The action of 
the iron alum should be watched under a microscope, and when 
the chromosomes of karyokinetic figures appear sharply defined, 
the slide should at once be placed in water and washed for at 
least an hour, since a very little of the iron alum, if left in the 
tissue, will cause the preparation to fade. If staining for details 
other than nuclei, the slide must be transferred to water as soon 
as the desired effect is produced. After the washing in water, 
the slide is passed through the alcohols, cleared in xylol, and 
mounted in balsam. This stain is excellent for the filamentous 
algae and fungi, and it keeps well in glycerine. For preparations 
to be mounted in balsam, erythrosin, fuchsin, or orange G make 
good contrast stains. Apply the second stain after the last 
washing in water. The second stain should always be very light. 

Chromosomes and centrosomes ("of those plants which have 
centrosomes") take a brilliant black, and other details, though 
not brightly colored, often show excellent definition. 

The times given above must not be accepted as final. Many 
prefer to wash in water for several hours after the first immersion 
in iron alum. A plan which has proved convenient and very 
successful is to put the slide into the iron alum in the morning, 
let it wash in water during the afternoon, leave it in the y 2 per 
cent, of hematoxylin over night, and finish the preparation the 
next morning. It is a long process, requiring care, patience, and 
judgment, but it is worth the effort. 

THE CARMINES. 

This group of stains, immensely popular a few years ago, has 
rapidly lost favor among botanists as newer stains and combina- 
tions have appeared. When it is desirable to stain in bulk, 
nothing has been found which will serve better than the car- 
mines. Only three of these stains will be considered : 



40 Methods in Plant Histology 

Greenacher's Borax Carmine. — 

Carmine, 3 g. 

Borax, 4 g. 

Distilled water, 100 cc. 

Dissolve the borax in water and add the carmine, which is 
quickly dissolved with the aid of gentle heat. Add 100 cc. of 
70 per cent, alcohol and filter. (Stirling.) 

The following is a slightly different method for making this 
stain from the ingredients mentioned above : Dissolve the 
borax in water, add the carmine, and heat gently for ten minutes ; 
after the solution cools, add the alcohol and filter; let the solu- 
tion stand for two or three weeks, then decant, and filter again. 

Stain the material in bulk from 50 per cent, alcohol one to 
three days, then treat with acid alcohol (50 cc. of 70 per cent, 
alcohol -f- 2 drops of hydrochloric acid) until the color becomes 
a clear red ; this may require only a few hours, but may take 
two or three days. The material may then be passed through 
the rest of the alcohols (six to twenty-four hours each), cleared, 
imbedded, and cut. After the sections are thoroughly fastened 
to the slide, the paraffin should be dissolved off with xylol. The 
balsam and cover may be added immediately, or the slide may 
be passed down through the alcohols for a contrast stain. 

Alum Carmine. — A 4 per cent, aqueous solution of ammonia 
alum is boiled twenty minutes with 1 per cent, of powdered 
carmine. Filter after it cools. (Lee.) 

Stain from water twelve to twenty-four hours and wash in 
water. No acid alcohol is needed, since the solution does not 
overstain. 

Alum Cochineal. — 

Powdered cochineal, 50 g. 

Alum, 5 g. 

Distilled water, 500 cc. 

Dissolve the alum in water, add the cochineal, and boil ; 
evaporate down to two-thirds of the original volume, and filter. 
Add a few drops of carbolic acid to prevent mold. (Stir- 
ling.) 



Stai?iing 4 1 

Stain as with alum carmine. A few years ago it was a very 
common practice to stain in bulk in alum cochineal and counter- 
stain on the slide with bismark brown. 

THE ANILINS. 

Many of the most brilliant and beautiful stains yet discovered 
belong to this group. These stains are so numerous that we 
shall not attempt to mention even their names, but shall con- 
sider only those which are in most common use by botanists. 
The following formula has proved to be fairly satisfactory for all 
the anilins mentioned in this account, but other formulae will be 
given for most of the stains : 

Make a 3 per cent, solution of anilin oil in distilled water ; 
shake well and frequently for a day; add enough alcohol to make 
the whole mixture about 20 per cent, alcohol ; add i.g. of cyanin, 
erythrosin, safranin, gentian violet, etc., to each 100 cc. of this 
solution. 

The anilins keep well in balsam, but not in glycerine. Xylol 
is a good clearing agent for all of them, but clove oil is very much 
better in case of gentian violet. Unfortunately, some of them 
do not give permanent stains. Some are acid, some basic, and 
some neutral. 

The rapidity with which sections must be transferred from 
one fluid to another makes many of them more difficult to man- 
age than the hematoxylins or the carmines, but the stains are so 
valuable that even the beginner should spend most of his time 
with the anilins. 

Many anilins stain quite deeply in one to twenty minutes, 
but if the stain washes out during the dehydrating process, stain 
longer, even ten to thirty hours, if necessary. If the stains are 
made up according to the formula mentioned above, transfer to 
the stain from 35 per cent, alcohol and from the stain to 35 per 
cent, alcohol, if the stain does not wash out too rapidly; if the 
stain washes out, try 50 per cent., 70 per cent., 85 per cent., 95 
per cent., or even directly to absolute alcohol. It will often be 
found impracticable to transfer from the stain to alcohols weaker 



42 Methods in Pla?it Histology 

than the 85 per cent., lest all the stain be washed out before the 
clearing agent is reached. 

Since the anilins are seldom used as single stains, but almost 
invariably in combination with other stains, the logical order 
will be disregarded and the stains will be treated, as they are 
used, in their most usual combinations. 

Cyanin and Erythrosin. — Stain in cyanin five to ten minutes 
or longer ; rinse quickly in alcohol, and then stain thirty seconds 
to one minute in erythrosin. If the cyanin washes out, stain for 
an hour, and if it still washes out, omit the rinsing in alcohol 
and transfer directly from the cyanin to the erythrosin. 

The erythrosin may be used first ; in this case stain for five 
minutes in erythrosin, transfer directly to cyanin, and stain for 
about ten seconds. Try 70 per cent, alcohol, but if the stains 
are lost, try 85 per cent., 95 per cent., or even transfer directly 
to 100 per cent. 

This is a good combination for general work, and if properly 
used is excellent for mitotic phenomena and the most delicate 
cytological work. Delafield's haematoxylin may be used with 
erythrosin. Stain first with the haematoxylin, and, after the pur- 
ple color has replaced the red due to the acid, stain lightly with 
erythrosin. Eosin may be used instead of erythrosin, but is less 
transparent. Eosin will be mentioned later in connection with 
special methods for algae and fungi. 

Flemming's Safranin-Gentian Violet-Orange. — Safranin has 
long been a famous stain for karyokinesis. This triple combina- 
tion was published in 189 1, but its value in plant cytology was 
not thoroughly appreciated until five or six years later, when its 
application was developed to a high degree of perfection by 
various investigators of the Bonn (Germany) school. 

According to Flemming, stain two to three days in safranin 
(dissolve 0.5 g. safranin in 50 cc. absolute alcohol, and after 
four days add 10 cc. distilled water); rinse quickly in water ; 
stain one to three hours in a 2 per cent, aqueous solution of 
gentian violet ; wash quickly in water, and then stain one to 
three minutes in a I per cent, aqueous solution of orange G. 






Staining 4 3 

Tranfer from the stain to absolute alcohol, clear in clove oil, and 
mount in balsam. 

The following method seems to be better for mitotic phe- 
nomena in plants : Transfer to safranin from 35 per cent, alcohol 
and stain sixteen to twenty-four hours. If the stain acts for 
only a few. hours, it washes out too rapidly to be controlled with 
any precision. (The safranin may be made up according to the 
formula given in the preceding paragraph or according to the 
general formula.) Rinse in water for a minute and then in 50 
per cent, alcohol until only nucleoli and the chromosomes of 
dividing nuclei retain the red color. If the alcohol does not 
wash out the stain sufficiently, dip the slide in acid alcohol (not 
more than 2 or 3 drops of hydrochloric acid to 100 cc. of 
50 per cent, alcohol). If acid has been used, wash for a 
moment in pure 50 per cent, alcohol, and then stain for four or 
five minutes in gentian violet (aqueous solution, or made up 
according to the general formula). Rinse for a few seconds in 
water and then stain about thirty seconds in a I per cent, aque- 
ous solution of orange G. Transfer from the stain directly to 
absolute alcohol and hasten the dehydrating by gently rinsing 
the slide in the fluid. One or two quick dips in the 95 per cent, 
alcohol before transferring to the absolute does not seem to do 
any harm and certainly saves the more expensive reagent. As a 
rule, not more than ten seconds can be allowed for dehydrating, 
because the gentian violet washes out so rapidly. The extreme 
transfer from the orange G, an aqueous stain, to absolute alco- 
hol, which must often be made to prevent the gentian violet 
from washing out, indicates that, for staining very thin sections 
on the slide, the series of alcohols from 35 per cent, up to 100 
per cent, is not at all necessary. 

Treat with clove oil for five to ten seconds. The clove oil 
not only clears, but it rapidly extracts the gentian violet, pro- 
ducing an elegant differentiation. Replace the clove oil by 
cedar oil, if you have strictly first-class cedar oil. Such cedar 
oil is very thin, light yellow in color, and has a faint odor of 
cedar wood. It costs about the same as clove oil. Do not get 



44 Methods in Plant Histology 

the expensive oil used for immersion lenses. If your cedar oil 
is colorless and has a strong odor, do not use it at all, but drain 
off the clove oil as thoroughly as possible by resting the edge of 
the slide on blotting paper for two or three minutes, and then 
mount in balsam. If any considerable amount of clove oil is 
left on the slide, the preparation is almost sure to fade. The 
cedar oil is not a solvent of gentian violet, and consequently 
there is little danger that the preparation will fade when this oil 
is used. Some investigators transfer from clove oil to xylol, 
but it seems to us that the brilliancy of the gentian violet is 
impaired. Chromosomes should take a clear red and the spindle 
fibers a bright violet. This combination is generally used after 
Flemming's solution, but seems to do equally well after other 
members of the chromic-acid series. Achromatic structures do 
not seem to stain well after corrosive sublimate or picric acid. 

Fuchsin. — Use a I or 2 per cent, solution in water or in 70 per 
cent, alcohol. Transfer to the alcoholic solution from 70 per 
cent, alcohol ; stain one to two hours ; differentiate the stain in 
I per cent, solution of picric acid in 70 per cent, alcohol — this 
may require thirty seconds or several minutes; rinse in 70 per 
cent, alcohol until a bright red replaces the yellowish color due 
to the acid, and then proceed as usual. 

Iodine Green. — A 1 per cent, solution in 70 per cent, alcohol 
is good for the vascular system of plants. This stain resists the 
washing-out process better than methyl green. Stain in iodine 
green at least an hour, and it is not a bad plan to stain over 
night ; rinse in 70 per cent, alcohol ; stain fifteen seconds to one 
minute in erythrosin, and proceed as usual. Methyl green may 
be made and used in the same way. 

Fuchsin and Iodine Green Mixtures. — Two solutions are kept 
separate, since they do not retain their efficiency long after they 

are mixed : 

0.1 g. fuchsin (acid). 
50 cc. distilled water. 
0.1 g. iodine green. 
50 cc. distilled water. 



Staining 4 5 

Jioo cc. absolute alcohol. 
1 cc. glacial acetic acid. 
0.1 g. iodine. 

Mix equal parts of A and B. Transfer to the stain from 
water. The proper time must be determined by experiment. 
Twenty-four hours might be recommended for a trial. Transfer 
from the stain directly to solution C and from C to xylol. 

Another formula : 

A. 0.5 g. acid fuchsin. 

B. 0.5 g. iodine green. 

Mix a pipette full of A with a pipette full of B ; stain two to 
eight minutes ; transfer to 85 per cent, or 95 per cent, alcohol, 
dehydrate rapidly, clear in xylol, and mount in balsam. Both 
these formulae are good for karyokinesis. 

Bismark Brown. — Use a 2 per cent, solution in 70 per cent. 
alcohol. If material has been stained in bulk in one of the 
carmines, a few minutes' stainino; on the slide with bismark brown 
gives a good contrast. It is particularly good for cell walls. 

Nigrosin. — Use a 1 or 2 per cent, solution in water. A few 
drops of this solution to a watch-glass full of water stains fila- 
mentous algae or fungi in one to three hours. The stain keeps 
well in glycerine or balsam, but it is hard to get these forms 
into balsam without more or less shrinking. 



CHAPTER VII. 

GENERAL REMARKS ON STAINING. 

It must be remembered that many things may be examined 
alive without killing, fixing, staining, or any of those processes. 
A filament of Spirogyra shows the chromatophore nicely if 
merely mounted in a drop of water; the nucleus may be visible, 
and the pyrenoids can usually be located. Of course, such a 
study is necessary if one is to understand anything about the 
plant, and in an elementary class this might be sufficient, but a 
drop of iodine solution applied to the edge of the cover would 
emphasize certain details, e.g., the starch in the pyrenoids would 
appear blue, the nucleus a light brown, and the cytoplasm a 
lighter brown. This illustrates at least one advantage to be 
gained by staining ; it enables us to see structures which would 
otherwise be invisible, or almost invisible. 

With so many stains at our disposal, it at once becomes a 
problem just which stain or combination to use in each particu- 
lar case. Beautiful and instructive preparations occasionally 
result from some happy chance, but uniform success demands 
skill and judgment in manipulation, and also a knowledge of the 
structures which are to be differentiated. Let us take a Vascular 
bundle for illustration. Safranin stains the xylem a bright red, 
but, with judicious washing, is entirely removed from the cam- 
bium and cellulose elements of the phloem. A careful staining 
with Delafield's haematoxylin now gives a rich purple color to 
the cellulose elements which were left unstained by the safranin, 
thus contrasting sharply with the lignified elements. If cyanin 
and erythrosin be used, the xylem takes the blue and the cam- 
bium and phloem take the red. Many terms have been given 
to indicate the affinity of certain tissues for certain stains. 
Auerbach used the terms "erythrophilous" and "cyanophilous " 
in 1890. This eminent zoologist studied spermatozoa and ova. 

47 



48 Methods in Plant Histology 

He found that, if preparations containing both spermatozoa and 
ova were stained with cyanin and erythrosin, the nuclei of the 
spermatozoa took the cyanin, while the nuclei of the ova pre- 
ferred the erythrosin; hence the terms " cyanophilous " and 
"erythrophilous." Auerbach regarded these differences as an 
indication of sexual differences in the cells. 

Rosen (1892) supported this theory, and even went so far 
as to regard the tube nucleus of the pollen grain as female, on 
account of its erythrophilous staining. In connection with this 
theory it was suggested that the ordinary vegetative nuclei are 
hermaphrodite, and that in the formation of a female germ 
nucleus the male elements are extruded, leaving only the ery- 
throphilous female elements ; and, similarly, in the formation of a 
male nucleus the female elements are extruded, leaving only the 
cyanophilous male elements. 

As long ago as 1884 Strasburger discovered that with a 
mixture of fuchsin and iodine green the generative nucleus of a 
pollen grain stains green, while the tube nucleus stains red. In 
1892 {Verhalten des Pollens) he discussed quite thoroughly the 
staining reactions of the nuclei. The nuclei of the small pro- 
thallial cells of gymnosperm microspores are cyanophilous like 
the male generative nuclei. The nuclei of a nucellus surround- 
ing an embryo-sac are also cyanophilous, while the nuclei of 
structures within the sac are erythrophilous. His conclusion is 
that the cyanophilous condition in both cases is due to poor 
nutrition, while the erythrophilous condition is due to abundant 
nutrition. A further fact in support of the theory is that the 
nuclei of the adventitious embryos which come from the nucellus 
of Funkia ovata are decidedly erythrophilous, while the nuclei 
of the nucellus to which they owe their food-supply are cyano- 
philous. 

In division stages nuclei are cyanophilous, but from anaphase 
to resting stage cytoplasm is taken into the nucleus, and the 
cyanophilous condition gradually changes to the erythrophilous. 

An additional fact in favor of this theory is that in Ephedra 
the tube nucleus which has very little cytoplasm about it is 



Ge?ieral Remarks o?i Stai?iing 49 

cyanophilous. Strasburger claims that there is no essential dif- 
ference between male and female generative nuclei, and subse- 
quent observation has shown that within the oospore the sex 
nuclei are alike in their reaction to stains. 

Malfatti (1891) and Lilienfeld (1892-3) claim that these 
reactions are dependent upon the amount of nucleic acid present 
in the structures. During mitosis the chromosomes consist of 
nearly pure nucleic acid and are intensely cyanophilous, but the 
protoplasm, which has little or no nucleic acid, is erythrophi- 
lous. There is a gradual transition from the cyanophilous con- 
dition to the erythrophilous, and vice versa, the acid structures 
taking basic stains and basic structures the acid stains. 

The terms "erythrophilous" and "cyanophilous" are falling 
into disuse, since the affinity is for basic and acid dyes, rather than 
for blue or red colors. That the terms are misnomers becomes 
evident when a combination like safranin (basic) and acid green 
(acid) is used, for the cyanophilous structures stain red, and the 
erythrophilous green. 

Probably but few investigators who have attained any pro- 
ficiency in microtechnique have not asked themselves how 
much dependence can be placed upon staining reactions as a 
means of analysis. Do staining reactions enable us to determine 
the chemical composition of a structure ? If two structures stain 
alike with Delafield's haematoxylin, does this mean that they 
have the same chemical composition ; or if, on the other hand, 
they stain differently, must they necessarily be different in their 
chemical composition ? Delafield's haematoxylin, when care- 
fully used, gives a rich purple color, but a careful examination 
will often show that in the same preparation some structures 
stain purple, while others stain red. Does this mean that the 
purple and red structures must have a different chemical com- 
position ? Many people believe that structures which stain dif- 
ferently with a given stain must be chemically different, but 
they readily agree that structures which stain alike are not 
necessarily similar in chemical composition. Chromosomes of 
dividing nuclei and lignified cell walls stain alike with safranin ; 



50 Methods in Plant Histology 

chromosomes and cellulose cell walls stain much alike with 
Delafield's haematoxylin ; but everyone recognizes that the 
chromosome is very different in its chemical composition from 
either the cellulose or the lignified wall. 

According to Fischer (1897 and 1900), stains indicate physi- 
cal but not chemical composition. Fischer experimented with 
substances of known chemical composition. Egg albumin was 
shaken until small granules were secured. These were fixed 
with the usual fixing agents, and then stained with Delafield's 
haematoxylin. The extremely small granules stained red, while 
the larger ones became purple. Since the granules are all alike 
in chemical composition, Fischer concluded that the difference 
in staining must be due to physical differences. With safranin, 
followed by gentian violet, the larger granules stain red and the 
smaller violet ; if, however, the gentian violet be used first, then 
treated with acid alcohol and followed by safranin, the larger 
granules take the red and the smaller the gentian violet. In 
root tips similar results were obtained. Safranin followed by 
gentian violet stained chromosomes red and spindle fibers violet, 
while gentian violet followed by safranin stained the chromo- 
somes violet and the spindle red. One often reads that chro- 
mosomes owe their strong staining capacity to nuclein, and 
especially to the phosphorus, but, according to Fischer, this is 
shown to be unfounded, since albumin gives similar results and 
yet contains no phosphorus, and is not chemically allied to 
nuclein. Delafield's haematoxylin is one of the so-called nuclear 
stains. The nuclei of animals and plants stain deeply with this 
reagent, but cellulose membranes, the dense protoplasm of 
embryonic cells, the pyrenoids of green algae, and many other 
structures resemble nuclei in their staining. The most critical 
work on this subject has been done by those who are investigat- 
ing the structure of the Cyanophyceae and Bacteria to determine 
whether these forms have nuclei or not. Butschli claims that 
the granules which stain red with haematoxylin are to be identi- 
fied with chromatin, while Fischer, whose results have just been 
given, claims that staining indicates merely physical differences. 



General Remarks on Staining 5 1 

The subject cannot yet be regarded as settled, but whatever may 
be true in regard to these conflicting theories, all agree that 
stains are of the highest importance in differentiating struc- 
tures, and in bringing out details which would otherwise be 
invisible. 



CHAPTER VIII. 

PRACTICAL HINTS ON STAINING. 

In later chapters specific directions will be given for making 
a series of preparations ranging from the lowest algae to the 
flowering plants, but a few suggestions will be made here. 

The number of stains in the catalogues is becoming so great 
that it is impossible to become proficient in the use of all of 
them. It is far better to master a few of the most valuable stains 
than to do indifferent work with many. The beginner, especially 
if rather unacquainted with the details of plant structure, may 
believe that he has an excellent preparation when it is really a 
bad, or at most an indifferent, one. To illustrate, let us suppose 
that a pollen mother-cell in a late spirem stage has been stained 
with cyanin and erythrosin. A preparation in which the cell 
merely shows a differentiation into nucleus and cytoplasm must 
be classed as bad ; if the nucleus shows a definitely outlined 
spirem thread, the preparation is better, but is still only indiffer- 
ent ; if the thread appears as a delicate red ribbon bordered by 
blue granules, the staining may be regarded as a success. If 
mitotic figures have been stained with cyanin and erythrosin, a 
first-class preparation should show blue chromosomes and red 
spindles ; if stained with safranin and gentian violet, the chromo- 
somes should be red and the spindles violet. 

In staining growing points, apical cells, young embryos, 
antheridia, archegonia, and many such things, the cell walls are 
the principal things to be differentiated, if the preparations are 
for morphological study. As a rule, it is better in such cases not 
to use double staining, but to select a stain which stains the cell 
walls deeply without obscuring them by staining starch, chloro- 
phyll, and other cell contents. For example, try the growing 
point of Equisetum The protoplasm of such growing points is 
very dense. If Delafield's haematoxylin and erythrosin be used, 

53 



54 Methods in Plant Histology 

the hematoxylin will stain the walls and nuclei, and will slightly 
affect the other cell contents, but the erythrosin will give the 
cytoplasm such a dense stain that the cell walls will be seriously 
obscured. It would be better to use hematoxylin alone. The 
same suggestion may well be observed in tracing the develop- 
ment of antheridia, archegonia, embryos, and similar structures. 
Permanent preparations are an absolute necessity for the 
greater part of most advanced work, but let us not imagine that 
we cannot examine anything until we have made a permanent 
mount. It would be impossible to make a permanent mount of 
the rotation of protoplasm. It is better for many purposes to 
look at motile spores while they are moving. Use Spirogyra 
while it is fresh and green (if you can) , and use permanent prep- 
arations only to bring out nuclei and other details which are 
not so easily seen in living material. Examples might be multi- 
plied. 



CHAPTER IX. 

THE CELLOIDIN METHOD. 

"Celloidin is a form of nitro-cellulose." It is very inflam- 
mable, but does not explode. It may be obtained in the form 
of tablets or cuttings, which have to be dissolved in a mixture 
of equal parts of absolute alcohol and ether. It is customary to 
use two solutions, a "thick" and a "thin." The thick solution 
(about 10 or 12 per cent.) should have about the consistency of 
thick syrup. The thin may be made by mixing equal parts of 
the thick and ether alcohol. 

As mentioned in the chapter on the "General Method," the 
killing, washing, and dehydrating are the same as for the paraffin 
method. After dehydrating in absolute alcohol the succeeding 
steps are as follows : 

1. Ether alcohol, 1 to 2 days. 

2. Thin celloidin, 2 to 6 days. 

3. Thick celloidin, 3 to 10 days. 

It seems better, however, to begin with about 2 per cent, 
celloidin and transfer successively through 4 per cent., 6 per 
cent., etc., to 12 per cent., or to allow the 2 per cent, to concen- 
trate by removing the cork for a short time each day. 

The material may now be imbedded and mounted upon 
a block at the same time. The blocks should have surface 
enough to accommodate the objects, and should be about one- 
fourth of an inch thick. White pine makes good blocks ; cork 
is much inferior. Place the block for a moment in ether alcohol 
and then dip into the 2 per cent, celloidin the end of the block 
which was left rough by the saw. With the forceps remove a 
piece of the material from the thick celloidin and place it upon 
the block, taking care to keep it right side up. Dip the block 
with its object first in thick celloidin, then in thin, and after 
exposing to the air for a few minutes drop it into chloroform, 

55 



56 Methods in Plant Histology 

where it should remain for about ten to twenty hours. It should 
then be placed in equal parts of glycerine and 95 per cent, 
alcohol, where it may be kept indefinitely. If the material is 
hard, like many woody stems, it will cut better after remaining 
in this mixture for a couple of weeks. 

In cutting, the knife should be set as obliquely as possible, 
and both the knife and the object should be kept wet with the 
mixture of glycerine and alcohol. As fast as they are cut the 
sections are transferred with a soft brush to 70 per cent, alcohol. 
The succeeding steps are the same as for free-hand sections, but 
many stains are not available because they stain the celloidin. 
Safranin and Delafield's haematoxylin, or Delafield's haematoxy- 
lin and eosin, are good combinations for celloidin sections. Do 
not use absolute alcohol for dehydrating, since it dissolves the 
celloidin, but transfer from 95 per cent, alcohol to Eycleshymer's 
clearing fluid (equal parts of bergamot oil, cedar oil, and car- 
bolic acid), which clears readily from 95 per cent, alcohol. 
Mount in balsam. 

The following schedules for staining celloidin sections will 
give the student a start. The times given will vary with the 
thickness of the section and character of the tissue. 

a. For staining in Delafield's haematoxylin and eosin : 

1 . Seventy per cent., 50 per cent., and 35 per cent, alcohol, 2 to 5 minutes 
each. 

2. Delafield's haematoxylin, 5 to 30 minutes. 

3. Wash in water, 5 minutes. 

4. Thirty-five per cent, and 50 per cent, alcohol, 2 to 5 minutes each. 

5. Acid alcohol (1 cc. hydrochloric acid+ 100 cc. of 70 per cent, alcohol) 
until the stain is extracted from the celloidin, or at least until the celloidin 
retains only a faint pinkish color. 

6. Seventy per cent, alcohol (not acid) until the purple color replaces the 
red due to the acid. 

7. Eosin (preferably a 1 per cent, solution in 70 per cent, alcohol), 2 to 5 
minutes. 

8. Eighty-five per cent., 95 per cent, alcohol, 2 to 5 minutes each. 
Remember that 100 per cent, alcohol is not to be used with celloidin 
sections. 



Celloidin Method 57 

g. Eycleshymer's clearing fluid until transparent, usually 1 or 2 minutes, 

but sometimes 5 or 10. 
10. Mount in balsam. 

b. For staining in safranin and Delafield's hematoxylin : 

1. Seventy per cent., 50 per cent., and 35 per cent, alcohol, 2 to 5 minutes 
each. 

2. Safranin, 6 to 24 hours, preferably the longer period. 

3. Acid alcohol (try about 1 drop of hydrochloric acid to 30 cc. of 70 per 
cent, alcohol) until the stain is removed from the celloidin or at least 
becomes very faint. 

4. Fifty per cent, and 35 per cent, alcohol, 2 to 5 minutes each. 

5. Delafield's hematoxylin, 2 to 5 minutes. 

6. Wash in water, 5 minutes. 

7. Thirty-five per cent, and 50 per cent, alcohol, 2 to 5 minutes each. 

8. Acid alcohol (the same as was used for the safranin) until the stain is 
extracted from the celloidin. If it is found that this double immersion 
in acid alcohol extracts too much safranin, the third step may be omitted 
or shortened by removing the sections to the 50 per cent, alcohol before 
the stain is thoroughly removed from the celloidin. 

q. Eighty-five per cent, and 95 per cent, alcohol, 2 to 5 minutes each. 

10. Eycleshymer's clearing fluid until cleared. 

1 1. Mount in balsam. 

The celloidin method has its disadvantages as well as its 
advantages. It is extremely slow and tedious, and it is rarely 
possible to cut sections thinner than io/x, while, on the other 
hand, it gives smoother sections. The entire absence of heat 
makes it very useful for delicate, succulent tissues. Stems and 
roots which cannot be handled at all in paraffin cut well in 
celloidin, and much larger sections can be cut than in paraffin. 

When material is to be imbedded, use celloidin as a last 
resort. Use paraffin when you can, celloidin when you must. 

I am indebted to my friend Mr. W. B. MacCallum for several 
suggestions in regard to this method. 



CHAPTER X, 



THE GLYCERINE METHOD. 



It is hard to get the filamentous algae and fungi into balsam 
without shrinking ; consequently, these forms are usually mounted 
in glycerine or glycerine jelly. 

Flemming's fluid and chromo -acetic acid are good fixing 
agents. Corrosive sublimate in water, or in 70 per cent, alcohol, 
used hot, is also to be recommended. For general morphology, 
stain for six hours or over night in a y 2 per cent, aqueous solution 
of eosin, transfer directly to a 1 per cent, solution of acetic acid 
in distilled water, and allow it to act for about five minutes ; wash 
thoroughly in water to remove the acid, and then put the 
material into a watch-glass in a 10 percent, solution of glycerine 
in water. The watch-glass should be kept as free from dust as 
possible, but should not be covered. As soon as the solution 
appears to be about as thick as pure glycerine, the material is 
ready for mounting. Place a small quantity of the material on 
a slide, arrange it carefully, add a small drop of glycerine, and 
a round cover. Seal with gold size (a varnish used by painters 
in laying gold leaf). None of the sealing media will stick to 
moist surfaces, hence it is essential that there should be only 
enough glycerine to come to the edge of the cover. If it is 
desired to mount rather large specimens, like the antheridia and 
oogonia of Chara, it is best to spin a ring on the slide, thus 
forming a shallow cell. Before the ring becomes hard the 
material may be mounted and sealed, or the ring may be allowed 
to harden, and just before a mount is to be made a very thin 
ring of the varnish may be added. The mount is firmer if the 
cover is not only sealed in the usual way, but also sticks to the 
ring underneath. 

If glycerine jelly is to be used, place the bottle in warm 

59 



60 Methods in Plant Histology 

water until the jelly becomes liquid, but avoid any unnecessary 
heat. Take the material from the glycerine, add a drop of the 
warm jelly, and seal as before. More detailed directions for 
mounting material in glycerine are given in the chapters on algae 
and fungi. 



Part II. 

In the preceding chapters the principles and methods of tech- 
nique have been described in a general way. It is often diffi- 
cult, especially for a beginner, to apply general principles to 
specific cases, and, besides, the types which he might select for 
the preparations might not form a symmetrical collection. Con- 
sequently, a series of forms has been selected which will not 
merely serve for practice in microscopical technique, but will 
also furnish the student with preparations for a fairly satisfactory 
study of plant structures from the algae up to the angiosperms. 
It is not at all our purpose to discuss general morphology, but 
rather to answer, by means of sketches and specific directions, 
the multitudinous questions which confront the instructor in the 
laboratory. For those who have had a thorough training in 
general morphology the following suggestions will be in some 
degree surperfluous. Those who are beginning the study of 
minute plant structure are referred to the standard text-books for 
descriptions of the plants mentioned here. 



CHAPTER XI. 



THALLOPHYTES. ALG-ffi. 
CYANOPHYCEJE. 
Wasserbliithe. — These forms occur as scums, often irides- 
cent, on the surface of stagnant or quiet water. Some of the 
commonest forms are Coelosphcerium and Anabcena {fig. ij). 
Some of the Chlorophycese also 
occur as Wasserbliithe. Where 
the material is very abundant, 
it may be collected by simply 
skimming it off with a wide- 
mouthed bottle, but where it is 
rather scarce, it is better to filter 
the water through a cloth, and 
finally rinse the algae off into a 
bottle. Enough formalin may 
now be added to the water in 
the bottle to make a 2 per cent, 
solution. The material may be 
kept here indefinitely, but after 
a few hours it is ready for use. 
If the forms are small, like 
Anab<z?ia, smear a slide lightly 

with Mayer's albumen fixative, paraffiD :sections stained in ***** and er y throsin - 
as if for paraffin sections, add a drop of the material and allow it 
to dry, heat the slide gently to coagulate the albumen, or immerse 
the slide in strong alcohol for a few minutes, and then proceed 
with the staining. Cyanin and erythrosin is a good combination 
for differentiating the granules. Delafield's hematoxylin, used 
alone, stains some granules purple and others red. Iron alum- 
haematoxylin is excellent for heterocysts. If the forms are large 
enough to collapse with such treatment, the glycerine method 
may be employed. 

63 




Fig. 



13- 



E 

Wasserbliithe. 



A, Ccelosphserium Kuetzingianum. B, Anabsena 

os-aquse. C, Anabsena gigantea. D and £, a 

heterocyst and spore of A. gigantea drawn from 



6 4 



Methods in Plant Histology 




If it is desirable to make paraffin sections, put the material, 
drop by drop, on a piece of blotting paper until an appreciable 
layer of sediment is obtained. Get the paper with its material 
into paraffin by the usual method, taking great care not to wash 
the algae off. After imbedding, trim away the 
paper and dip the block in melted paraffin. 
Sections can now be cut and stained in the 
usual manner. 

Oscillaria. — For most purposes it is best to 
study Oscillaria in the living condition. It is 
readily found in watering troughs, in stagnant 
water, on damp earth, and in other habitats. 
The commonest forms have a deep blue-green 
fig i Oscillaria or Drowmsn color. For the purposes of identi- 
Portions of two fiia- fication and herbarium specimens, the material 

merits, the one at the . . . r . 

right showing a hormo- may simply be placed on a slip or mica and 

gonium, h. J r J r r 

allowed to dry. When wanted for use, add a 
drop of water and a cover, and the mount is ready for examina- 
tion. For sections or 
for glycerine mounts 
fix in chromo-acetic 
acid. {Fig- 14.) 

Rivularia.- — This 
form is readily found 
on the underside of the 
leaves of water-lilies 
[Nuphar, Nymphcea, 
etc.) , but is also abun- 
dant on submerged 
leaves and stems of 
other plants. It oc- 
curs in 

translucent, gelatinous nodules of various sizes. Chromo-acetic 
acid gives beautiful preparations, but good results can also be 
secured from formalin or picric-acid material. 

The most instructive preparations for morphological study 




Rivulark 



A, nodule crushed under cover-glass. B, four filaments more 
the form Of highly magnified, showing heterocysts at the base. 



Thallophytes. Algcz 




Fig. 16. Tolypothrix. 

b, a false branch, h, hetero- 
cysts. 



can be obtained by the glycerine method. 
Stain in eosin or Mayer's haem-alum. When 
ready for mounting, crush a small nodule 
by pressing on the cover-glass. Fig. ij is 
drawn from such a preparation. The par- 
affin method is easily applied, since the 
gelatinous matrix keeps the plants in 
place. Glceotricliia, Nostoc, and forms of 
similar habit may be prepared in the same 
way. 

Tolypothrix. — This form occurs as small 
tufts, either floating in stagnant water or 
attached to plants and stones. It furnishes 
an excellent example of false branching. 
{Fig. 1 6.) Scyto?iema is a similar form 
which is fairly common. The glycerine 
method should be employed for perma- 
nent preparations, but this, like all small 
filamentous algae, may be dried on mica for herbarium pur- 
poses. 

CHLOROPHYCEiE. 

The ponds, ditches, and rivers of any locality will yield an 
abundance and variety both of the unicellular and multicel- 
lular members of this group. The unicellular and filamentous 
members, together with such forms as Volvox, are best prepared 
by the glycerine method. The structure is so much more com- 
plicated than in the Cyanophyceae that it demands far more care 
and skill to make good preparations. Chromo-acetic acid is a 
good killing and fixing agent for the whole group, but Flem- 
ming's fluid (weaker solution) seems to be a little better in some 
instances. Very good results have been obtained by adding 
about 5 cc. of I per cent, osmic acid to ioo cc. of chromo- 
acetic acid (Schaffner's formula). A formula which gives satis- 
factory results with Spirogyra may cause plasmolvsis with 
Cladophora. The given filament should be placed under the 
microscope in the fixing agent, and, if plasmolysis occurs, the 



66 



Methods i?i Plant Histology 





B 



Fig. 17. Vaucheria. 

A , Vaucheria geminata. B, V. sessilis. 
0, oogonia. 



a, antheridia. 



chromic should be weakened or the acetic strengthened until the 
suitable proportions are determined. This is a slow process, but 
Cladophora and Vauclieria are almost sure to shrink without it. 
About twenty-four hours in any of the chromic series and a four 
to ten hours washing in water will be sufficient for members of 

this group. Only a few 
o of the most familiar forms 
will be mentioned. 

Vaucheria. — This form 
can always be obtained in 
greenhouses, especially in 
the fernery, where it forms 
a green felt on the pots. 
The greenhouse form is 
likely to be Vaucheria sessilis. Another species, V. geminata, is 
very common in the spring, when it may be found in ponds and 
ditches. {Fig. 17.) It is extremely difficult to get mounts show- 
ing the nuclei. The following method is sometimes successful : 

Chromoacetic acid (Schaffner's formula), 24 hours. 

Wash in water, 4 to 10 hours. 

Iron alum, 2 to 4 hours. 

Water, 1 5 to 30 minutes. 

One-half per cent, hematoxylin, over night. 

Wash in water, 5 to 10 minutes. 

Iron alum until details become clear. This may take only a few minutes, 

but may take an hour. 

Wash thoroughly in water, 1 to 4 hours. 

Ten per cent, glycerine and allow the glycerine to thicken. 

Mount and seal. 



Cladophora. — This is found attached to sticks and stones in 
quiet or running water. It is easily recognized by its character- 
istic branching. {Fig. 18.) The nuclei of the ccenocytic seg- 
ments are readily brought out by the method just described for 
Vaucheria. Alum carmine and Mayer's haem-alum are also good 
stains for the nuclei. 

Hydrodictyon. — This is popularly known as the "water-net." 
Nets of all sizes should be selected for study. The segments 



Thallophytes. Algce 



6 7 



are coenocytic, and the nuclei are hard to differentiate except in 
the younger segments. The method given for Cladoplwra yields 
good results. The young nets forming within the older seg- 
ments are easily demonstrated by Mayer's haem-alum. Paraffin 
sections stained in cyanin and erythrosin, iron 
alum-haematoxylin, or the safranin-gentian violet- 
orange combination will repay the trouble, especially 
in case of the oldest segments within which new 
nets are forming. The habit is beautifully shown 
in preparations stained with eosin. The eosin ( 1 per 
cent, aqueous solution) should act for about twenty- 
four hours. Then transfer to 1 per cent, acetic 
acid for a few minutes. If the stain comes out 
rapidly in the acid, one minute may be sufficient, 
but if the stain does not wash out, it is better to 
let the acid act for four or five minutes. Wash 
thoroughly in water to remove all trace of acid, 

or the preparation will 
fade. Transfer to 10 per 
cent, glycerine and pro- Fixed in chromo . 

1 1 / »t" \ acetic acid, stained in 

Ceed aS USUal. [Plg.IQ.) Haidenhain-s iron 
_ . r^. . . alum-hsematoxylin. 

Spirogyra. — This is 
probably the most widely known of all 
the algae, and, fortunately, it is rather 
easy to obtain beautiful and instructive 
preparations. The following is a good 
fixing agent for most Spirogyras: 

Chromic acid, -j^- g. 

Glacial acetic acid, -fo g. 

Water, 99 cc. 

The addition of 1 cc. of 1 per cent, osmic acid seems to 
improve it without causing any blackening. Flemming's weaker 
solution is excellent. If it causes too much blackening, as it 
probably will, the material, after being washed, should be placed 
in weak peroxide of hydrogen (one part H 2 2 to three parts 
H 2 0) until the blackening due to osmic acid disappears. After 





Fig. 18. Cladophora 



Fig. 19. Hydrodictyon. 
A small portion of a young net. 



68 



Methods in Plant Histology 



a moment's washing in water it is then ready for the stain. The 
iron alum-haematoxylin, as just described, brings out the nuclei 
and pyrenoids with great distinctness. {Fig. 20.) A few min- 
utes in aqueous eosin after the last washing in water often gives 
a beautiful differentiation, but the preparations will be quite 




Fig. 20. Spirogyra 



From material fixed in chromo-acetic acid and stained in iron alum-haematoxylin. A, 
single cell showing nucleus, chromatophore, and pyrenoids-. B, a nucleus undergoing 
division. C, a resting nucleus. D, zygospores, each showing two nuclei. 

inferior if the eosin is allowed to stain too deeply. Mayer's 
hoem-alum is a better stain for stages in conjugation. 

It is difficult to get Spirogyra into paraffin without shrinking, 
but it can be done. Watch carefully and note where plasmolysis 



Thallophytes. Alga 



69 



occurs. There will probably be little or no trouble until 
the transfer from 100 per cent, alcohol to the clearing agent. 
Make this transfer as gradual as may be necessary. After the 
pure xylol or other clearing agent is reached, add a lump of 
paraffin large enough to saturate the clearing fluid at a tempera- 
ture of 40 to 45 C. Allow the xylol to evaporate at this 
temperature and imbed as usual, taking care to 
keep the filaments as nearly parallel as possible. 
Many elegant combinations, like cyanin and 
erythrosin, fuchsin and iodine green, safranin- 
gentian violet-orange, and others not available 
for glycerine preparations, can be used with 
paraffin sections. It is comparatively easy to 
get any such alga into celloidin. Safranin and 
Delafield's haematoxylin then make a good 
combination. 

Zygnema — Use the same methods as for 
Spirogyra. In staining conjugating material the 
stain should be extracted until the four chroma- 
tophores of the zygospore become distinct. The 
nuclei are comparatively small and unsatisfac- 

-T-., , 11 , 1 ,i 11 The filament at the left 

tory. Ine stellate chromatopnores are well shows three zygospores and 

1 . .. . x r • i one parthenogenetic spore 

brought out by alum carmine, ii iron alum- which is distinguished by 

having only two chromato- 

haematoxylin and eosin are used, the eosin may phores. The filament on the 

J J right shows two cells, each 

well be much deeper than in case of Spirogyra. with a pair of stellate chro- 

i- c °- / matophores. Drawn from 

( FtP" 2 T \ material fixed in 2 per cent. 

V o" ') formalin and stained in iron 

Diatoms.— Diatoms and desmids have been ^-^ at °*y lin - 
variously classified, and their position is not yet fully determined. 
Living diatoms are often found clinging in great numbers to fila- 
mentous algae, or forming gelatinous masses on various submerged 
plants. It is difficult to get really good preparations showing the 
nucleus and chromatophores. If the diatoms are clinging to fila- 
mentous algae, the algae with the diatoms attached may be put 
into chromo-acetic acid (twenty-four hours), washed in water, 
stained, passed up through the alcohols, and cleared in xylol, 
or, better, in clove oil or bergamot oil, which do not dry up so 




70 



Methods in Plant Histology 






rapidly. Here the diatoms may be picked or scraped off from 
the other algae, which will probably have become much shrunken 
by this treatment. Mount in balsam. Haidenhain's iron alum- 
haematoxylin is recommended for the nucleus and the centro- 
some, which is quite prominent in diatoms. Delafield's haema- 
toxylin and erythrosin give a good view of the nucleus and 

chromatophore. If a glycer- 
ine mount is preferred, the 
iron alum-haematoxylin is a 
good stain. 

When the material is in 
gelatinous masses, it may be 
fixed in chromo-acetic acid 
and imbedded in paraffin. 
There will, of course, be some 
difficulty in cutting, and many 
frustules will be broken, but 
there will, nevertheless, be 
occasional views which show 
details better than when the 
diatoms are mounted whole. 
The silicious shells of dia- 
toms are among the most beautiful objects which could be exam- 
ined with the microscope {fig. 22) . To obtain perfectly clean 
mounts requires considerable time and patience, but when the 
material is once cleaned, preparations may be made at any time 
with very little trouble. Diatom enthusiasts have devised numer- 
ous methods for cleaning diatoms, and separating the various 
forms from each other, but we shall give here only a few simple, 
practical methods. 

Material for mounts of frustules of living forms "may be 
obtained by skimming off the brownish scum found on ponds, 
by squeezing out water weeds, by scraping sticks and stones 
which are covered at high water, or from the mud of filter beds 
at pumping works, or in other places. The material is put in a 
dish of water, and after it has settled the water is decanted. 





Diatoms. 



255- 



A, Pleurosigma angulatum. B, Navicula dactylis. 
C, Synedra biceps. D, Gomphonema sphaerophorum. 
E, Triceratium sp. 



Thallophytes. Algce 71 

This is repeated until the water will clear in about one-half hour. 
The sediment is then treated with an equal bulk of sulphuric 
acid, after which bichromate of potash is added until all action 
ceases. After a couple of hours the acid is washed out. To 
separate the diatoms, place the sediment in a glass dish with 
water, and when the water becomes clear give the dish a slight 
rotary motion. This will bring the diatoms to the top, when 
they may be removed with a pipette and placed in alcohol. To 
mount, place a number in distilled water, evaporate a few drops 
of the mixture on a cover-glass, which is then mounted on a 
slide in balsam." (From a review of Dr. Wood's paper on 
14 Diatoms," Jour. App. Mic, March, 1899.) 

Many scouring soaps and silver polishes contain large quan- 
tities of fossil diatoms, and the diatomaceous earths are particu- 
larly rich. Break up a small lump of such material and boil it 
in hydrochloric acid. A test-tube is very convenient for this 
process. Let the diatoms settle, pour off the acid, and then 
wash in water. As soon as the diatoms settle, the water should 
be poured off. The washing should be continued until the 
hydrochloric acid has been removed. When the washing is 
complete, pour on a little absolute alcohol, and after a few 
minutes pour off the alcohol and add equal parts of turpentine 
and carbolic acid. The material will keep indefinitely in this 
condition and may be mounted in balsam at any time. In mak- 
ing a mount, put a little of the material on a slide and allow it 
to become dry, or nearly dry, and then add the balsam and cover. 
If the balsam should be added too soon, the diatoms are likely 
to move to the edge of the cover. 

Desmids. — When these forms are very abundant, they may be 
treated like the filamentous algae, except that extreme care must 
be taken lest the desmids be lost while changing fluids. It 
often happens that a single desirable desmid appears when 
examining field collections. In such a case, remove it with a 
fine pipette, and get it into a drop of water on a clean slide, 
invert it over a bottle of 1 per cent, osmic acid for a minute, 
leave the slide exposed to the air until the water has almost all 



72 



Methods i?i Plant Histology 



evaporated, and then add a drop of 10 per cent, glycerine. In 
a few hours (six to twenty-four) put on a cover and seal. It 
requires more time, care, and patience than it is worth to 
attempt staining in such a case. [Fig. 23.) 

Oedogonium. — In selecting material it will be better for teach- 
ing purposes to choose the larger monoecious forms. The nuclei, 

pyrenoids, and chromatophores are 
easily differentiated. Mayer's haem- 
alum is a good stain, especially for 
the antheridia. Alum carmine or 
eosin will bring out the " caps." 
{Fig. 24.) 

Chara. — This form 
is so large and coarse 
that it hardly pays to 
mount it in glycerine. 
If a glycerine mount is Q 
desired to show the an- 
theridia and oogonia in 
position, spin a ring of 
cement on the slide, 
thus making a cell in 
which small portions of the plant may be mounted. 
For paraffin sections select the tip of the plant, 
a piece about half an inch in length. Sections of 
this may show, not only the large apical cell, but 
also various stages in the development of antheri- 
dia and oogonia. Delafield's haematoxylin is a 

. r 1 ■ 1 11 1 r 1 a > antheridia. o, 

very good stain tor the apical cell and tor the oogonium. Drawn 

from material fixed in 

development of antheridia and oogonia. The later 1 per cent, chromic 

1 ° acid, and stained in 

stages in the development of antherozoids are Mayer's hsem-aium. 
brought out more clearly by the safranin-gentian violet-orange, 
or by cyanin and erythrosin, but here unusual care must be taken 
not to stain too deeply. 

Good preparations showing shield, manubrium, capitula, and 
filaments may be obtained by staining in bulk in alum carmine 




Fig. 23. Desmids. X 255. 
From glycerine preparations. Not stained 
A, Cosmarium pectinoides. B, Closte- 
rium cucumis. C, Staurastrumcornutum 
£>, Arthrodesmus octocornis. 




Fig. 24. Oedogonium 
nodulosum. 



Thallophytes. Algce 



73 



and then crushing the antheridium under the cover-glass after 
the specimen is in balsam. [Fig* 25 •) 

PKJEOPH.YCEJE. 

The brown algae are almost exclusively marine. The slime, 
so prevalent in the group, often makes the technique difficult. 

Ectocarpus. — Fix in 
chromo-acetic acid 
(twenty-four hours), 
wash in fresh water, 
since the salt of sea 
water may cause incon- 
venience in subsequent 
processes. Stock mate- 
rial should be passed up 





Fig. 



Chara. 



A, portion of a branch showing an antheridium, a, and an 
oogonium, o. B, median longitudinal section of an apical cell. 
Drawn from a preparation fixed in chromo-acetic acid, and 
stained in Delafield's hcematoxylin. 

to 70 per cent, alcohol for safe keeping. 
Eosin or Mayer's haem-alum are good for 
glycerine mounts. If paraffin sections are 
to be made, the material must be brought 
very gradually from absolute alcohol into 
the clearing agent, and from the clearing 
agent into the paraffin. {Fig. 26.) 

Other filamentous members of the 
group, as well as the more delicate mem- 
branous forms, may be treated like Ecto- 
carpus. 

Fucus. — Fucus may be fixed, washed, and preserved like 
Ectocarpus. It is difficult to get paraffin sections across the whole 
fertile branch, but elegant sections may be obtained by cutting 



Fig. 26. Ectocarpus confervoides. 

From a preparation stained in 
Mayer's hsem-alum, and mounted in 
glycerine. X 255. m, multilocu- 
lar sporangium, u, unilocular spo- 
rangium. 



74 



Methods in Plant Histology 



narrow strips containing a few conceptacles. The safranin- 
gentian violet-orange combination is good for such sections. For 
such views as are represented in fig. 2j, C and D, the material 

should be stained 




Fig. 27. 



D 

Fucus vesiculosus. 



A, small portion of plant showing bladders and fruiting branches. 
One -half natural size. B, transverse section of fruiting branch showing 
oogonial^ conceptacles. X 6. C, antheridia and paraphyses. From a 
preparation fixed in chromo-acetic acid, stained in borax carmine, teased 
out and mounted in balsam. X 255. D, oogonium showing five of the 
eight oospheres. Prepared as in C. 



in bulk in borax 
carmine or alum 
carmine. The 
process for borax 
carmine is as fol- 
lows : 

1. Borax carmine, 24 
hours. 

2. Acid alcohol (2 
drops of HC1 in 
50 cc. of 70 per 
cent, alcohol), until 
the color becomes 
a clear red. This 
may take an hour 
or even a day. 

3. Seventy to 100 per 
cent, alcohol, 2 
hours each. 

4. Clear in cedar oil, 
bergamot oil, or 
oil of cloves. 

5. Tease out the con- 
tents of the con- 
ceptacles suffi- 
ciently to show 
details, and mount 
in balsam. 



The process for alum carmine is the same, except that no 
acid alcohol is used. 

Sections like that shown in B are easily cut in celloidin. 
After staining in borax carmine or alum carmine, imbed in 
celloidin in the usual way. After hardening the celloidin in 
chloroform, put the block into 95 per cent, alcohol for two or 
three hours, and then into Eycleshymer's clearing fluid (equal 
parts bergamot oil, cedar oil, and carbolic acid), until thoroughly 
cleared. The block may be left here indefinitely, and sections 



Thallophytes. Algce 



75 



may at any time be mounted in balsam as soon as they are 
cut. 

Chorda, Lami?iaria, and similar forms may be treated like Fucus. 



B 




RHODOPHYCE.E. 

The red algae belong almost exclusively to salt water, but a 
few genera are found only in fresh water, usually in running water, 
and a few forms occur both ^ 
in salt and in fresh water. 

The technique is more 
difficult than in the case of 
the brown algae. Until some- 
thing better is suggested, the p 
same method of fixing and 
washing may be used as for 
the brown algae. Picric acid, 
corrosive sublimate, and ab- t 
solute alcohol have been 
tried, but do not give as good 
results as the chromo-acetic * 
acid or Flemming's fluid. 

Batrachospermum. — This 
is a green, fresh-water mem- 
ber of the red algae. It is 
not very uncommon in small 

Streams (Fip~ 28 \ From a preparation stained in Mayer's hsem-alum and 

V * 'J mounted in glycerine. A, portion of plant showing 

TVi^ rpllc orp CA email branches and several cystocarps. X 25. B, a procarpic 

IIIC CClib die bU bllldll branch showing carpogonium (/) , and trichogyne (/), 

.1 , • , • 1 11 ,1 1 '1 with an antherozoid (s) attached. X 255. C, a younger 

mat It IS naraly WOrtn Wniie branch showing carpogonium and trichogyne. X 255. 

, . . , D, branch with three antherozoids. X 255. 

to attempt sectioning them. 

Very good preparations showing the nuclei may be obtained by 
staining in Mayer's haem-alum, or Haidenhain's iron alum-haema- 
toxylin. After the material is in glycerine ready for mounting, 
tease out a small portion, and still further dissociate the filaments 
by tapping smartly on the cover. 

Material stained in eosin shows the external structure well, 
but may not bring out the nuclei. 





% W9 



Fig. 28. Batrachospermum moniliforme. 



7 6 



Methods in Plant Histology 



Polysiphonia. — For preparations like those shown in fig. 29 
eosin is a very good stain. To get a brilliant coloring, stain 

for about twenty-four hours, 
so that the 1 per cent, acetic 
acid may be allowed to act 
for several minutes without 
making the stain look weak. 
Wash thoroughly in water. 
Not the slightest trace of 
color should be allowed to 
come out in the glycerine. 

Sections showing the cen- 
tral and peripheral siphons 
and other gross features are 
easily cut in celloidin. It is 
not very difficult to cut par- 

Fig. 29. Polysiphonia fibrillosa. J l 

From a preparation fixed in chromo-acetic acid, stained airm Sections, DUt tne nUCiei 
in eosin, and mounted in glycerine. X 255. A, an 11 j 1 » , 

antheridium. B, a cystocarp with carpospores. C, are SO Small and. SO narQ tO 

tetrasporic branch with tetraspores. , , , 

bring out that such prepa- 
rations had better be left for the specialist. 




CHAPTER XII. 



THALLOPHYTES. FUNGI. 









K\,^-^r' 



9V 



SCHIZOMYCETES. 

Bacteria. — The methods of modern bacteriological technique 
are so numerous and so specialized that we must refer to laboratory 
manuals for instruction in this subject. The method given here 
will merely enable the student to study 
the form and size of those bacteria 
which are more easily demonstrated. 

Foul water at the outlets of sewers 
and such places will usually afford an 
abundance of coccus, bacillus, spiril- 
lum, and beggiatoa. forms. Place a 
drop of the water on a slide, heat it 
gently until the water evaporates, then 
stain with fuchsin or methyl violet, 
dehydrate, clear in xylol, and mount 
in balsam {fig. jo). 

Fine preparations may be obtained 
by inoculating a mouse with A?ithrax 
or some other form, and then cutting 
paraffin sections of favorable organs. 
Gentian violet with a faint bismark 
brown for a background makes a good 
combination. The following schedule gives good results with 
Anthrax and many other bacteria: 

i. Gentian violet, 5 minutes. 

2. Rinse in water a few seconds. 

3. Gram's solution (iodine 1 gram, potassium iodide 2 grams, water 300 cc.) 
until the color is almost or quite black ; this will generally require 1 or 
2 minutes. 

4. Ninety-five per cent, alcohol until the color has nearly disappeared. 

77 




*»> 



? " C 



Fig. 30. Bacteria, X 535. 

A , Bacillus anthracis, from a paraffin 
section cut from the liver of a mouse. 
Fixed in chromo-acetic acid, stained in 
methyl violet and bismark brown, and 
mounted in balsam. B, Staphylococ- 
cus pyogenes aureus. From a prepa- 
ration stained in gentian violet. C. 
Spirillum sp. From a preparation 
stained in fuchsin. 



78 



Methods in Plarit Histology 



5. Rinse in water and examine. If the bacteria are well stained, a counter- 
stain for the background may be added. 

6. Erythrosin, 3 or 4 seconds ; or bismark brown 5 or 10 seconds. 

7. Ninety-five and 100 per cent, alcohol, dehydrating as rapidly as possible. 
Not more than 5 or 10 seconds can usually be allowed. 

8. Xylol. 

9. Balsam. 

Leptothrix may often be obtained by scraping the inside of 
the cheek. Beggiatoa, a form with oscillating movements like 
Oscillaria, is often found in foul water. Its presence may be 
indicated by whitish patches on the bottom. 

It is doubtful whether the bacteria possess even a morpho- 
logical forerunner of the nucleus of higher plants, consequently 
there need be no disappointment if the larger forms, like some 
of the Beggiatoas, fail to show a nucleus. 

MYXOMYCETES. 

With the exception of a few forms like Fnligo (often found 
on oak stumps and on oak bark in tan yards), the Myxomycetes 
are small, and are usually overlooked by collectors. (-Fig.ji.) A 

careful examination of rotting logs 
in moist woods will usually reveal 
an abundance of these delicate 
and beautiful organisms. They 
should be pinned to the bottom 
of a box for safe carrying. For 
herbarium specimens they are 
simply allowed to dry, and are 
then fastened with glue or paste 
to the bottom of a small box. 

Plasmodia and young sporan- 
gia may be fixed in chromo-acetic 
acid or Flemming's fluid. Sections. are easily cut in paraffin, 
and should not be more than 5 \x in thickness, and should be 
thinner, if possible. Acid fuchsin and iodine green is a good 
stain. Delafield's hematoxylin used alone or with a little 
orange G. is also to be recommended. Excellent methods for 




Fig. 31 



c 

Myxomycetes. 



Growing on rotten wood. A , Hemitrichia rubi 
formis. X 20. B, Stemonitis ferruginea. Nat 
ural size. C, Trichia varia. X t.V 2 . 



Thallophytes. Fungi 



79 



living cultures were given in the January and February (1898) 
numbers of the Journal of Applied Microscopy . 

PHYCOMYCETES. 

Mucor. — This familiar mold appears with great regularity on 
bread. The following is a sure and rapid method for obtaining 
Mucor: Place a glass tumbler in a plate of water, put on the 
tumbler a slice of bread which 
has been exposed to the air for a 
day, and cover with a glass jar. 

To obtain such a series as is 
shown in A-D of fig. 32, the ma- 
terial should be studied before 
the sporangia begin to turn black. 
The phase in the life-history indi- 
cated in F-H is rarely seen. The 
writer would be glad to hear from 
any who have met this phase, 
especially if the information could 
be accompanied by a few zygo- 
spores. 

Corrosive sublimate (4 per 




Mucor stolonifer. X 255. 



A-D, successive stages in the development 
of trie sporangium. Drawn from living material. 
£, columella with a few spores adhering. F-H. 
Cent.) in 50 P er Cent. alCOhol, stages in the formation of the zygospore. From 

a preparation fixed in corrosive sublimate, 
USed hot, may be recommended stained in Delafield's hsematoxylin, and mounted 
J in glycerine. 

as a fixing agent. Haem-alum, or 

Delafield's haematoxylin, is good for glycerine preparations. 
Do not stain too deeply. A very satisfactory study may be 
made from the living material. 

Cystopus. — This fungus is quite common on Cruciferae, where 
the white "blisters" or "white rust" form quite conspicuous 
patches. 

Affected portions of leaves and stems should be fixed in 
chromo-acetic acid and cut in paraffin. Sections 5 ^ or less in 
thickness will be found most satisfactory. Safranin-gentian 
violet-orange seems to be the best stain for differentiating the 
nuclei. {Fig.jj.) 



8o 



Methods in Plant Histology 




It is more difficult to get good sec- 
tions of the plant in the oosporic condi- 
tion. The oosporic phase of Cystopus bliti 
is easily recognized on Amara?itus, where 
the oospores may be seen with the naked 
eye by holding the leaf up to the light. 
While better nuclear staining can be se- 
cured with chromic or Flemming mate- 
rial, it will be found somewhat easier to 
cut material which has been fixed in pic- 
ric acid (i per cent, solution in 70 per 
cent, alcohol). Celloidin sections, stained 
in Delafield's hematoxylin, can be rec- 
ommended for showing the position of 

Fig. 33. Cystopus candidus .. . , , i ' • i • i,, i i 

on Capseiia. oogonia and antnendia, although such sec- 

w ra v s " « c V of a bl!ster °V he tions are too thick to give satisfactory views 

Jear. X 255. b rom a preparation o J 

fixed in Flemming's fluid and f j_u niiHpi 
stained in safranin-gentian vio- Ui Lilc UU ^ 1C1, 
let-orange. 

ASCOMYCETES. 

This group, popularly known as the "sac fungi," contains an 
immense number of saprophytic and parasitic forms. Yeast, 
green mold on cheese and leather, leaf curl of peach, black 
knot of cherry and plum, and the -powdery mildews are familiar 
to everyone. The few objects selected will enable the student 
to experiment, but he must not be discouraged if success does 
not crown the first attempt, for the group presents many diffi- 
culties. 

Saccharomyces. — Until somewhat recently it was considered 
rather difficult to demonstrate the nucleus of the yeast cell. 
With fresh growing yeast the following method by Wager 
should be successful : Fix in a saturated aqueous solution of 
corrosive sublimate for at least twelve hours. Wash success- 
ively in water, 30 per cent, alcohol, 70 per cent, alcohol, and 
methyl alcohol. Place a few drops of alcohol containing the 
cells on a cover, and when nearly dry add a drop of water. 
After the yeast cells settle, drain off the water and allow the 






Thallophytes. Fungi 



81 



cells to dry up completely. Place the cover, or slide, with its 
layer of cells in water for a few seconds, and then stain with a 
mixture of fuchsin and methyl green, or fuchsin and methylen 
blue. Mount in glycerine or in balsam. 

Eurotium. — For class use or for permanent preparations it is 
best to select nO/0?-^ r\ 

rather y o u n ~ ' 
material which 
shows various 
stages in devel- 
opment, from the 
swollen end of the 
hypha to the ripe 
spore (fig. 34) . 




A B 



C 

Fig. 34 



XJ JOUR APP MIC 
Eurotium. 



From material growing on a hectograph pad. Fixed in chromo-acetic 
acid, stained ineosin, and mounted in glycerine. A -E, successive stages in 

The nuclei are development, x 375. 

exceedingly small, and can hardly be demonstrated with eosin. 

Iron alum-haematoxylin would be better. 

The following schedule will give good mounts for habit study: 

1. Fix in a saturated solution of corrosive sublimate in 50 per cent, alcohol. 
The addition of 1 cc. of glacial acetic acid to 100 cc. of this solution 
improves it. Use it hot. 

2. After it cools, transfer to 50 per cent, alcohol and add, a few drops at a 
time, the iodine solution which is used in testing for starch. At first the 
brownish color caused by the iodine will disappear, but after a certain 
amount has been added the brown color will remain. The material is 
then ready for the next step. 

3. Thirty-five per cent, alcohol, 5 minutes. 

4. Eosin (aqueous), at least 5 minutes ; a day will do no harm. 

5. Put the material into 1 per cent, acetic acid for 15 seconds to 2 minutes. 
The material should still have a vivid red color when taken from the acid. 

6. Wash in water. Use a considerable volume of water or change the water 
several times. If the acid is not all washed out, the preparations will fade. 

7. Ten per cent, glycerine, and allow the glycerine to thicken. 

8. Mount in glycerine or glycerine jelly, and seal. 

Any of the filamentous fungi — like Mucor, Thamnidium, 
Peroiwspora, Penicillium, Pythium, Saprolegnia, etc. — can be 
mounted in this way. Saprolegnia, however, is much more satis- 
factorv if stained in iron alum-haematoxylin. 



82 



Methods i?i Plant Histology 



A very rapid method for the unicellular and filamentous 
forms may be added : 

1. One hundred per cent, alcohol, 2 minutes. 

2. Eosin (aqueous), 2 minutes. 

3. One per cent, acetic acid, 2 to 10 seconds. 

4. Wash in water 5 minutes, changing frequently. 

5. Mount directly in 50 per cent, glycerine, and seal. 

If the material gets through the first four stages without 
shrinking, but collapses at the fifth, put it into 10 per cent. 

glycerine and allow it to 
thicken as usual. 

Uncinula necator. — 
The mildews are found 
throughout the summer 
and autumn on the leaves 
of various plants. The 
lilacmildew( Microsphcera 
aim) and the mildew on 
the Virginia creeper 
(Uncinula necator) are 
particularly abundant. 
For herbarium purposes 
they may be preserved 
by simply drying the 
leaves under light pres- 
sure. When mounted for 
examination, the leaf 

Fig. 35. A, Uncinula necator on Ampelopsis quinquefolia. SUOUld be SOaked in Water 

X 192. Four asci containing ascospores have been forced out r r • r, 

by pressing out the cover. Fixed in hot corrosive sublimate, IOT a lew minutes, alter 
stained in fuchsin, and mounted in balsam. B, a conidiospore ; . . . 

and C, an appendage of Microsphsera alni, drawn from living which the peritiieCia may 
material. X 192. r J 

be scraped off and 
mounted in water. The asci may then be forced out by press- 
ing smartly on the cover. {Fig. J5.) 

For permanent mounts of entire perithecia with appendages, 
fix in 3 per cent, formalin twenty-four hours, wash in water one 
hour, stain in aqueous eosin twenty-four hours, treat with 1 per 




JOUR APR MIC. 



^C 



Thallopliytes. Fwigi 



83 



cent, acetic acid one minute, wash thoroughly in water, and then 
transfer to 10 per cent, glycerine, which should be allowed to 
concentrate as usual. If chromic acid, corrosive sublimate, or 
alcohol be used for fixing, the appendages become brittle, and 
very easily break off. However, the chromo-acetic mixtures 
are better if it is desired to make paraffin sections showing the 
development of the perithecium with its asci and spores. For 
this purpose the omnipresent 
Erysiphe commune on Polygonum 
aviculare is exceptionally favor- 
able, because, after the material 
is fixed and in alcohol, the whole 
mycelium, with the developing 
perithecia, may be stripped from 
the leaf without the slightest 
difficulty, thus avoiding the 
necessity of cutting the leaf in 
order to get the fungus. The 
safranin- gentian violet -orange 
combination seems to give the 
best results, although cyanin 
and erythrosin are quite satis- 
factory when the stains are P eT } the /} a 

J stained in 

properly balanced. 

YTr1/ii>in T\/T £ 1'1 several days in equal parts of g5 per cent, alcohol 

Ayiaria. Many tOrmS, like and glycerine, and then imbedded in celloidin. Not 




JOURAPPMIC. 

Fig. 36. Xylaria. 

A , transverse section of a young stroma showing 
X 8. Fixed in chromo-acetic acid, 
bulk in alum carmine, imbedded in celloi- 
din, and mounted in balsam. B, two asci with 
spores. X 245. The mature stroma was soaked for 



stained. 



Xylaria, Ustilina, Hypo xy Ion, and 
Nummularia, in their mature condition, are woody and so 
extremely brittle that it is almost impossible to cut them. As 
good a plan as any seems to be to cut sections of the stroma 
about one-eighth of an inch thick, soak them in equal parts of 
glycerine and 95 per cent, alcohol, and then imbed them in 
celloidin in the usual way. They might be cut without imbed- 
ding, but most of the asci and spores would then be lost. 

The younger stages, showing the development of perithecia 
and asci, are more interesting, and can be cut in paraffin and 
stained with ease {fig. J 6) . 



8 4 



Methods in Plcrnt Histology 



Peziza. — The Pezizas and related forms are fleshy, and pre- 
sent but little difficulty in fixing, cutting, or staining. They are 
abundant in moist places, on decaying wood, 
or on the ground. The apothecia have the 
form of little cups, which are sometimes 
black and sometimes flesh -colored, but 
often orange, red, or green. For the devel- 
opment of ascospores in the ascus, Flem- 
ming's fluid (weaker solution), followed by 
safranin-gentian violet-orange, has given the 
best results with thin sections where the 
mitotic figures are to be studied. Cyanin 
and erythrosin is also to be recommended. 
Such sections should not be more than 
5 ft in thickness. For a general morpho- 
logical preparation, such as is shown in 
the figure, it is better to stain in bulk in 
alum carmine or in Delafield's hematoxy- 
lin, and then tease out the asci in gly- 
cerine or balsam. Sections thick enough 
to show the entire ascus are not usually 
as satisfactory as such teased preparations. 
{Fig- 37-) 




Fig. 37. Peziza odorata. 
Three asci and many para- 
physes. X 245. Fixed in cor- 
rosive sublimate, stained in 
bulk in alum carmine. Teased 
out and mounted in balsam. 



^CIDIOMYCETES. 

The ^Ecidiomycetes comprise the rusts (Uredinea) and the 
smuts ( Ustilaginece ) . 

Puccinia graminis. — The common rusts of wheat and oats 
are familiar to everyone. The uredospores, or summer spores, 
known as the red rust, and the teleutospores (last spores), or 
winter spores, known as the black rust, are found in unfortunate 
adundance, but the aecidium stage on the barberry is not neces- 
sary for the vigorous development of rust in the United States, 
and is seldom found. Most teachers are obliged to depend upon 
botanical supply companies for this material. There are, how- 
ever, various aecidia which are as good, or even better, for 



Thallophytes. Fungi 



85 



morphological study. The aecidia growing on Euphorbia maculata 
(spotted spurge), and on Ariscema triphy ilum (Jack-in-the-pulpit) 
are much easier to cut, and seem easier to stain. Delafield's 
haematoxylin, followed by a very light stain in erythrosin, is 




ii/OVHAPBmC.. 

Fig. 38. Puccinia graminis. 

A, transverse section of barberry leaf showing aecidia and spermagonia. X 7. B, longitudinal 
section of a single aecidium. X 192. Fixed in Flemming's weaker solution and stained in Delafield's 
haematoxylin. C, a single spermagonium. X 192. Fixed and stained as in B. D, three uredospores 
growing on oats. X 375. Fixed in 2 per cent, formalin, stained in bulk in alum carmine, and teased 
out in glycerine. E, section of young teleutospores on oats. X 375. Fixed in picro-acetic acid and 
stained in cyanin and erythrosin. G, F, H, three ripe teleutospores from a leaf of oats showing varia- 
tion in form. X 375. A germinating teleutospores. X 375. 



86 Methods in Plant Histology 

good for both secidia and spermagonia, especially after Flem- 
ming's fluid. It is rather difficult to get good sections of uredo- 
spores and teleutospores, because the leaves of wheat and oats 
are refractory objects to cut. The cutting is easier after picro- 
acetic acid than after corrosive sublimate or the chromic-acid 
series. (Fig. 38.) 

Every class which studies the rusts should attempt to germi- 
nate the uredospores and teleutospores. For this purpose the 
hanging drop culture may be employed. Cement a rubber or 
zinc ring to the slide, or simply smear the lower surface of the 
ring with vaseline and press it tightly against the slide ; smear 

the upper surface of the ring 



Fig. 39. 



with vaseline, and over it invert 
I the cover-glass with a shallow 
drop of water containing the 
spores (Jig- J<?). The uredo- 
spores germinate readily all summer, but it is said that the 
teleutospores will germinate only in the spring following their 
maturity. However, the teleutospores of many species, like 
Puccinia xanthii on Xa?ithium canadense (cocklebur), will germi- 
nate as soon as they ripen and will serve equally well for study. 
If a particularly good specimen is secured, it may be preserved 
by the method previously described for desmids, except that in 
this case it might be worth while to attempt staining with Mayer's 
hsem-alum, or with eosin. 

The smuts may be studied in the living material. The fol- 
lowing method, recently described by Ellis, is worth remember- 
ing : A supply of smutted barley may be obtained by sowing 
soaked, skinned barley that has been plentifully covered by 
Ustilago spores. In such material it is easy to trace stages in 
the development of spores. Free-hand sections of ears about 
three-eighths of an inch long show the mycelium and spore clus- 
ters. If smutted ears be removed and kept floating on the 
water, the spores continue to develop and often germinate. For 
paraffin sections desirable stages should be fixed in Flemming's 
fluid or picro-acetic acid. Delafield's hematoxylin, followed by 




Thallophytes. Fimgi 87 

a very light touch of erythrosin or acid fuchsin, will give a good 
stain. 

For a study of the germinating spores and conidia, cultures 
may be made in beerwort on the slide or in watch crystals. 
Harper's method of making preparations from such material is 
ingenious and will undoubtedly prove valuable in making mounts 
of various small plant and animal forms. A drop of the material 
is taken up with a capillary tube and is then 
gently blown out into a drop of Flemming's 
weaker solution (fifteen minutes or an hour was 
sufficient for the fungus spores). Cover a 
slide with albumen fixative, as if for sections.) 
A drop of the material, without previous wash- 
ing, is drawn up into the capillary tube and 
touched lightly and quickly to the surface of 

a ■ r 1 , 1 _ r \ JOUKAPR.MC. 

the albumen. A series or such drops, almost j s 

as small as the stippled dots in a drawing, may S 

be applied to the slide. The fixing agent may FlG>4 °- Coprinus ; comatus - 

Transverse section of a 

now be allowed to evaporate somewhat, but portion of one of the giiis 

1 showing a part of the trama, 

the preparation must not be allowed to dry. '» t nd . s ^ v f ral basi . dia - $> 

1 J- J each with four stengmata, 

As the slide is passed rapidly through the £ om Sp °^ y h T ^hfSnV 
alcohols, the albumen is coagulated, and the mata ' X75 °' 
preparation may be treated just as if one were dealing with 

ribbons of sections. 

BASIDIOMYCETES. 

This is an immense group, of which the mushrooms, toad- 
stools, puffballs, and bracket fungi are the most widely known 
representatives. 

Coprinus comatus. — This is the common shaggy-mane mush- 
room. Cut from the cap pieces about one-fourth of an inch 
square, and fix in chromo-acetic acid or in Flemming's fluid. Por- 
tions in which the gills have just begun to turn brown will show the 
spores still attached to the sterigmata ( fig. 40) . If the gills have 
become dark brown or black, the spores will wash off before the 
sections can be mounted. Look in portions in which the gills 
are still white or only slightly changing color for the develop- 



88 Methods in Plant Histology 

ment of basidia and spores. The nuclei, although rather small, 
are brought out nicely by safranin-gentian violet-orange. The 
same procedure may be observed for other forms of similar con- 
sistency, like many members of the genera Boletus, Hydnum, 
Polyporus, Lycoperdon, etc. Leathery or woody forms like Stereum 
and many species of Polyporus had better be fixed in picro-acetic 
acid and imbedded in celloidin. Young stages of Cyathus or 
Crucibulum (bird's-nest fungi) cut easily in paraffin, but the 
older stages cut much better in celloidin. It is hard to get the 
very soft, watery forms like Tremella into paraffin without shrink- 
ing, but sections as thin as 10 i^ may be cut in celloidin. While 
this is too thick to give satisfactory views of such small nuclei, 
it brings out very clearly the general morphological structures. 

THE LICHENS. 

The lichens are usually regarded as difficult forms. In 
younger stages they occasion no trouble, but an old apothecium 
or a leathery thallus often fails to cut well. Difficulties may be 
minimized by using prolonged periods. The following schedule 
has proved satisfactory for the thalli and mature apothecia of 
Physcia, Us?iea, Sticta, Collema, Parmelia, and Peltigera: 

1. Chromo-acetic acid (medium solution, p. 28), 2 to 4 days. 

2. Wash in water, 6 to 24 hours. 

3. Thirty-five, 50, 70, 85, and 95 per cent, alcohols, 6 to 24 hours each. 

4. One hundred per cent, alcohol, 2 to 4 days, changing 2 or 3 times. 

5. Mixtures of alcohol and xylol, 1 to 2 days. 

6. Pure xylol, 6 to 24 hours. 

7. Xylol and paraffin on the bath, 1 to 2 days. 

8. Paraffin at 54 to 6o°, changing once or twice, 3 to 6 days. 

9. Imbed in as thin cakes as possible. 

Cyanin and erythrosin is a very good stain for lichens. The 
algae stain blue and the filaments of the fungus take the red. 
Where the association of the alga and the fungus is rather loose, 
as in Dichonema, more satisfactory mounts can be made by stain- 
ing in eosin, or hsem-alum and eosin, and then teasing slightly 
with needles and mounting in glycerine. 



CHAPTER XIII. 

BRYOPHYTES. 

The Bryophytes, comprising the two groups Liverworts 
{Hepaticce) and Mosses (Musci) , present a great diversity of 
structure, some being so delicate that good preparations are very 
uncertain, while others are so hard that it is difficult to get satis- 
factory sections. Between these extremes, however, there are 
many forms which readily yield beautiful and instructive prepa- 
rations. 

If but one fixing agent should be suggested for the entire 
group, it would be chromo-acetic acid with ^ g. chromic acid 
and y 2 cc acetic acid to ioo cc. of water. It should be allowed 
to act for at least twenty-four hours, and probably two or three 
days would be better. Always make an effort to get the mate- 
rial into paraffin, using celloidin only as a last resort for refrac- 
tory structures which resist infiltration and for very delicate 
structures which persist in collapsing. As one gains in experi- 
ence and carefulness, the number of cases which seem to demand 
celloidin will become fewer and fewer. 

Instead of treating forms in a taxonomic sequence, we shall 
consider first the gametophyte structures under the headings 
thallus, antheridia, and archegonia, and shall then turn our atten- 
tion to the sporophyte. 

HEPATICiE. 

Some of the liverworts are floating aquatics, but most of 
them grow on logs or rocks or upon damp ground. They are 
found at their best in damp, shady places. Many of them may 
be kept indefinitely in the greenhouse. Riccia, Ricciocarpus, 
Marchantia, Conocephalus, Asterella, and many others vegetate 
luxuriously, and often fruit if kept on moist soil in a shady part 
of the greenhouse, and they do fairly well in the ordinary labo- 
ratory if covered with glass and protected from too intense 



90 



Metlwds in Plant Histology 




A. L> Joufl.flpp.Mic 

Fig. 41. Ptilidium ciliare. X 420. 
A, longitudinal. B, transverse section of the apex of the 
leafy gametophyte. Fixed in Flemming's weaker solution, 
stained in a mixture of acid fuchsin and iodine green. Ten 
microns. 



light. The living plants are very desirable, since they not only 
furnish the best possible material for habit work and the coarser 
microscopic study, but they also enable one to secure complete 
series in the development of the various organs. 

The Thallus. — In many cases it will not be necessary to make 
a special preparation for the study of the thallus, since prepara- 
tions of antheridia, archegonia, or sporophytes may include good 

sections of vegetative por- 
tions. This is particularly 
true of forms like Riccia, 
where the various organs 
are not raised above the 
thallus. In forms like Mar- 
chajitia, where the antheri- 
dia, archegonia, and. sporo- 
phytes are borne upon 
stalked receptacles, it is 
better to make separate preparations to show the structure of 
the mature thallus. Sections intended to show the structure 
of the mature thallus should be 15 \x to 25 /jl in thickness, but 
sections to show the growing point and development of the 
thallus should not be thicker than 10 ft. Material showing apical 
cells and development of the thallus is easily gotten into paraf- 
fin, even in forms like Ricciocarpus, which in their mature condi- 
tion are in danger of collapsing. The apical region of the foliose 
Jnngerma?i?iiace(E [fig. 41 s ) affords an excellent opportunity for 
studying the development of the plant body from a single apical 
cell. If mixtures containing osmic acid are used for fixing, 
there may be difficulty in the staining, even after using per- 
oxide of hydrogen. Corrosive sublimate-acetic, Carnoy's fluid, 
or chromo-acetic acid are better for apical regions. A fairly 
vigorous staining with a mixture of acid fuchsin and iodine 
green often brings the walls out very sharply. Chromo-acetic 
.acid, followed by Delafield's haematoxylin or bismark brown, is 
good for the apical cells and developing regions, but a light 
counter-stain with erythrosin improves preparations of the 



Bryophytes 



91 



mature thallus. After corrosive sublimate-acetic the material 
may be stained in bulk with alum cochineal or alum carmine, 
thus giving fairly good preparations and saving considerable 
labor. 

Antheridia. — If you have the material, it is not difficult to 
get good preparations showing the development of antheridia. 
In forms like Asterella, Pcllia, etc., cut out a small portion of the 
thallus bearing the antheri- 
dia. The piece should not 
be more than a quarter of 
an inch square, and if it can 
be smaller, so much the ^— <— r T 
better. For early stages of 2/m^J"a 
the antheridia of Marchan- 






Fig. 42. Asterella hemisphaerica. X 255. 

Successive stages in the development of antheridia. 
Fixed in chromo-acetic, stained with Delafield's ha:ma- 
toxylin. Section 10 microns thick. 



tia select young antheridio- 
phores which still lie close 
to the thallus. These 
readily cut as thin as 5 fi, 
and a single slide will usu- 
ally show a more complete 
series than is represented in the figure of Asterella (Jig. 42), but 
after the stalk begins to lengthen, the younger stages become 
infrequent, and it is not always easy to cut thin sections. Dela- 
field's hematoxylin or bismark brown serves very well for such 
stages as are shown in the figure. The protoplasm of the young 
antheridia is so dense that the addition of a counter-stain is 
almost sure to injure the preparation by obscuring the cell walls. 
For stages older than that represented in D, showing the devel- 
opment of the spermatozoid, the paraffin must be rather hard 
(melting at 55 C. to 65 C.) , and the sections should not be 
thicker than 5 fi, while 2 \x or 3 /jl is best. For such stages use 
the safranin- gentian violet -orange combination, Haidenhain's 
iron alum-haematoxylin with or without a faint trace of eryth- 
rosin or orange G, or use a mixture of acid fuchsin and methyl 
green. Nothing but practice and patience will bring success in 
such critical work. 



9 2 



Methods i?i Plant Histology 



If antherozoids are found escaping, transfer them to a small 
drop of water on a clean slide, invert the drop over a I per cent, 
solution of osmic acid for two or three minutes, allow the drop 
to dry up, pass the slide through the flame two or three times, 
as in mounting bacteria, and then stain sharply in acid fuchsin. 

This should show the 
general form of the an- 
therozoid, and will usu- 
ally bring out the cilia. 

The Archegonia. — The 
methods for archegonia 
are practically the same 
as for antheridia. Too 
much stress cannot be 
laid upon the importance 
of carefully selecting the 
material. Use very small 
pieces, and, before pla- 
JiM.fypAk D cm g them in the fixing 

Fig. 43. Marchantia polymorpha. X 400. - . • ,1 , 1 

■ agent, trim them to such 

A, three early stages in the development of the archegonia. 

Delafield's hsematoxylin. B, young archegonium showing two- a. shape that the position 
neck canal cells and the central cell before the cutting off of " r 

the ventral canal cell. Fuchsin and methyl green. C, mature ^f 4-V.^ arnVif±crr\r\ic\ will Kp> 

archegonium just ready for fertilization. Safranin-gentian 0t "ie arOiegOnia Will De 

violet-orange. D, young embryo. Delafield's hsematoxylin. accura telv knOWn even 

after the pieces are imbedded in paraffin. For stages like_/z£\ 43, 
A and B, Delafield's hsematoxylin is a good stain, and 10 /m is 
about the right thickness. For stages like C, in such forms as 
Marchantia, where the necks are long and often somewhat curved, 
it is better for general purposes to use sections from 1 5 \x to 
20 fJi in thickness. If it is desired to obtain preparations show- 
ing the cutting off of the ventral canal cell, the development of 
the oosphere, and the process of fertilization, the sections should 
be from 5 ft to 10 ^ in thickness, and the same staining may be 
used as for the development of antherozoids. For archegonia 
containing young embryos, like that shown in D, Delafield's 
hematoxylin without any counter-stain gives beautiful prepara- 
tions when the staining is well done. It is easier for the beginner 




Bryophytes 



93 



to get good preparations with the safranin-gentian violet-orange 
combination. 

The Sporophyte. — Sporophytes in early stages of develop- 
ment often yield good preparations without very much trouble, 
but in later stages they are frequently difficult to cut on account 
of the secondary thickening of the capsule 
wall and the stubborn extine of the mature 
spores. It is hard to get Ricciocarpus into 
paraffin without shrinking, and the same 
thing may be said of other forms which 
have such loose tissue with large air cavi- 
ties. For stages like that shown \r\ftg. 44, 
as well as for older sporophytes, it will be 
found more uniformly satisfactory to use 
celloidin, and cut the sections from 20 ^ 
to 30 fJL thick. 

For a study of archegonia, antheridia, 
young sporophytes, and also for the devel- 
opment of the thallus, it is better, even in 




Fig. 



JouaflppAls 
Ricciocarpus natans. 




JwilV-^C 



Fig. 45. Pellia epiphylla. 



A , habit sketch of sporophyte. X 10. B, small portion of 
sporophyte (at X of A), showing the capsule wall, the spores, and 
the elaters. Fixed in chromo-acetic acid and stained in cyanin 
and erythrosin. Ten microns. 



Young sporophyte inclosed in 
the archegonium. Spore mother- 
cell stage. All the cells of the 
sporophyte except a single peri- 
pheral layer (dotted in the figure) 
produce spores. Fixed in picro- 
acetic acid and stained in Dela- 
field's haematoxylin. Celloidin 
section 30 microns in thickness. 



forms like Ricciocarpus, 
to use paraffin. Pro- 
longed fixing in chro- 
mo-acetic acid (two to 
six days), thorough 
dehydrating, and 
gradual transfer from 
alcohol to xylol, and 
from xylol to paraffin, 



and also a moderate temperature in the bath (not more than 
50° C), will often bring the material through in fine condi- 
tion. 



94 



Methods in Pla?it Histology 



Forms like Pellia cut well in paraffin, especially in younger 
stages, but even in case of mature sporophytes it is not neces- 
sary to resort to celloidin. In Pellia and Co?iocephalus the spores 
are very large and have a rather thin wall. Both these genera 
show a peculiar, intrasporal development of the gametophyte, 
i. e., the gametophyte develops to a considerable extent before 
it ruptures the spore wall. ( Fig. 45.) For the older sporophytes 
of Marchantia it is better not to cut the whole receptacle, but 




Jour./1pp.M/<l 



Fig. 46. Anthoceros laevis. 



A, longitudinal section of lower portion of sporophyte imbedded in the gametophyte. X 45. B, 
transverse section of lower portion of sporophyte. X 200. Delafield's hsematoxylin. Ten microns. 
C, vegetative cell from lower portion of the sporophyte. X 560. Fixed in Flemming's weaker solution 
and stained in a mixture of acid fuchsin and iodine green. Five microns. D, spore mother-cell showing 
three of the four chloroplasts with numerous starch grains. The nucleus is in the metaphase of the first 
division. X 560. Fixed in Flemming's weaker solution, stained in safranin-gentian violet-orange. 
Five microns. 

rather to remove the branches so that they may be cut separately. 
For the very best preparations of mature sporophytes it will pay 
to trim away the gametophyte structures, leaving only enough to 
show the foot with a few of the surrounding cells. Sections 5 ft 
to 10 fi thick can be made without much difficulty from material 
prepared in this way. 

Among the Bryophytes no form affords a better oppor- 
tunity for studying the development of spores than Anthoceros, 



Bryophytes g 5 

since a single longitudinal section of the sporophyte may show 
all stages, from earliest archesporium to mature spores {fig. 46) . 
For studies like A and B, chromo-acetic material cut 10 ^ thick 
and stained in Delafield's hematoxylin is very good. The 
starch grains in the chloroplasts take a beautiful violet color 
with the safranin-gentian violet-orange combination. It is very 
difficult, however, to bring out the details of nucleus or chloro- 
plast on account of the minute size of these structures. The 
drawings from which C and D were reproduced were made with 
a one-sixteenth oil immersion objective. The drawings, like all 
the others illustrating the Bryophytes, were reduced one-half by 
photography. 



CHAPTER XIV. 



BRYOPHYTES. 

MUSCI. 

Material for a study of the mosses is much more abundant, 
and a series of stages in the development of the various organs 
is easily secured ; but it is much more difficult to obtain good 
preparations, because so many of the 
structures are hard to cut. Chromo- 
acetic acid is to be recommended as 
the most satisfactory fixing agent, but 
where structures are refractory and very 
likely to make trouble in cutting, it will 
often be found more satisfactory to use 
picro- acetic acid in the 70 per cent, 
alcohol, since material fixed in this 
reagent does not become as hard or as 
brittle as that fixed in any of the chro- 
mic-acid series. 

Antheridia. — It is easy to find mate- 
rial for a study of antheridia, because, 
in so many cases, the antheridial plants Twenty mi 

U J a- 4. J 4. vu 4. diaofPoly 

can be detected at once without even a 
pocket lens. Fwiaria, with its bunch of antheridia as large as a 
pin-head, is extremely common everywhere. Spring is the best 
time to collect it, but it is found fruiting in the autumn and 
sometimes in summer; besides, it is easily kept in the green- 
house, where it may fruit at any time. Bryum proliferum has a 
still larger cluster of antheridia, which may be seen at a distance 
of several yards. Polytrichum also has a large cluster of anthe- 
ridia surrounded by reddish leaves, so that the whole is some- 
times called the moss "flower." In making preparations of 
Polytrichum these colored leaves should be carefully removed 

97 




Fig. 47. 



A, archegonia of Webbera candi- 
cans. X 104. Celloidin section, 
microns. B, young antheri- 
olytrichum commune. X 420. 



9 8 



Methods in Plant Histology 



after the material has been gotten into 70 per cent, alcohol. A 
single antheridial plant of Polytrichum often furnishes a fairly 
complete series of stages in the development of antheridia. 
{Fig. 47.) I n a ^ cases the stem should be cut off close up to 
the antheridia, for many of the moss stems cut like wire. It is 
not necessary to use celloidin for antheridia, nor is it desirable, 

except where sec- 
tions from 20 /jl to 
50 \i thick are want- 
ed for habit work. 
Delafield's haema- 
toxylin is recom- 
mended for staining. 
Archegonia. — 
Since the necks of 
the archegonia are 
usually long and 
more or less curved, 
it is necessary, for 
habit work, to cut 
sections as thick as 
20 ft or 30 fJL in or- 
der to get a view of 
JoM.flpp.M»& an archegonium in a 

Fig. 48. Funaria hygrometrica. i , • /~> i 

4 >s . single section. Cel- 

A, apex of young sporophyte showing endothecium and amphithe- 

cium. X 420. Chromo-acetic acid and Delafield's hsematoxylin. Ten loidin IXiaV be USed 
microns, B, C, and D, transverse sections of a sporophyte of the same •> 

age as A, taken at three different levels. X 255. Ten microns. for SUCh DTeDara- 

tions, but for the development of the archegonium, the oosphere, 
the canal cells, and also for the process of fertilization, it is better 
to use paraffin. For the thick celloidin sections the material 
may be stained in bulk in alum cochineal, but thin paraffin sec- 
tions should be stained on the slide with more critical stains. 

(^r- 47) 

The Sporophyte. — It is often difficult to get good mounts of 
sporophytes. In the younger stages the calyptras are likely to 
interfere with cutting, while in the older stages the peristome, or 




Bryophytes 



99 



hard wall of the capsule, occasions the trouble. If an attempt is 
made to remove the calyptra in young stages, like A of fig. 48, 
the apex of the sporophyte usually comes with it. While 
picro-acetic acid material cuts more easily, chromo-acetic acid 
followed by Delafield's haematoxylin gives so much sharper dif- 
ferentiation in stages like those shown in^. 48 that it is better 




JOUH [{pft.Mlt. 



Fig. 49. Funaria hygrometrica. X 420. 

A, longitudinal section of capsule. B, transverse section of capsule of about the same age as A. 
The columella, archesporium, outer spore case, two layers of chlorophyll-bearing cells, and the beginning 
of the air spaces can be distinguished at this stage. Delafield's haematoxylin and erythrosin. Ten 
microns. 

to use harder paraffin (55 to 6o° C.) and make an effort to get 
preparations from chromic material. 

Stages like that shown in fig. 4Q are cut with comparative 
ease, for the calyptra is easily removed, and the capsule wall is 
not yet hard enough to occasion any difficulty. The cell walls 
are so easily stained in moss capsules that a light counter-stain 
with erythrosin or acid fuchsin may be used to bring out the 
L.ofC. 



100 



Methods in Plant Histology 




cytoplasm and plastids without appreciably obscuring the cell 
walls. Funaria and Bryum afford an excellent study in the 
development of the capsule, since all the structures of a highly 
differentiated moss sporophyte are present, and Bryum is par- 
ticularly easy to cut in stages like those 
shown in fig. jo. 

Sporophytes, in their more mature 
stages, are almost sure to present con- 
siderable difficulty in cutting. For gen- 
eral work fairly good preparations may 
be gotten from celloidin material, but it 
is worth while to try paraffin, for it is 
sometimes successful, and when it does 
succeed it is far 
JovRfafyc* superior. As soon 
fig. 5 o. Bryum. x 200. as the cell walls 

Portion of a nearly mature capsule K^m-in f/~> fViiVU<=>tn 

showing operculum, annulus, peris- Degin IO tUlCKen, 
tome, and three cells of the sporo- • .1 H^w»1nr» 

genous tissue. Fixed in Flemming's ab m LIie UCVClUp- 
weaker solution, stained in safranin . r .1 

and Delafield's hsematoxylin. Fif- ment OI me peri- 
teen microns. r 

stome, sarranin is 
an excellent stain, and this, followed by 
Delafield's hematoxylin, will give an ele- 
gant differentiation in the older stages of 
the sporophyte. After capsules have be- 
gun to turn brown it will be almost impos- 
sible to infiltrate them unless they are 
pricked with a needle. 

The mature sporophytes of Sphagnum 
[fig. 5/) are exceptionally hard to cut. It 
will be worth while to prick the capsule 

1 L Longitudinal section of mature 

with a needle when the material is col- sporophyte, showing also the upper 

portion of the pseudopodium and 

lected. This will allow the fixing agent ^ e iJj£3& SZSgfi 
to penetrate readily, and will also facilitate Paraffin - Tenmicrons - 
the infiltration of paraffin or celloidin. The puncture causes 
only a slight damage, and need not reach the really valuable 
portion which is to furnish the median longitudinal sections. 




Fig. 51. Sphagnum. X 24. 



Bryophytes 1 1 

Protonema and teased mounts of antheridia and archegonia 
may be made directly in 50 per cent, glycerine without fixing or 
staining. While this method is often recommended, we have 
found it better to use 10 per cent, glycerine and allow it to con- 
centrate. Mounts made in this way retain their green color for 
a long time. 



CHAPTER XV. 

PTERIDOPHYTES. 

This group, including the Filicineae, Equisetinese, and Lyco- 
podinese, or, more popularly, the ferns, horsetail rushes, and 
club mosses, is familiar to everyone. Material is abundant, 
and so easily recognized that anyone who pays a little attention 
to collecting can, in a single season, get a fine supply for a study 
of the group. Some desirable forms may not be present in all 
localities, but these will be few and can be obtained at a reason- 
able price from those who make a business of collecting. 

FILICINEJE. 

Without attempting to follow any taxonomic sequence, the 
methods of preparing the various structures of the homosporous 
forms will be presented, and then the peculiarities of the hetero- 
sporous members will be considered. 

The Prothallia. — Ripe spores of some fern or other can be 
obtained at any greenhouse at any time in the year, and spores 
of most of our native ferns germinate well and produce good 
prothallia, even if the sowing is not made for several months 
after the spores have been gathered. 

Fine prothallia of Pteris aquilina have been grown two years 
after the spores were gathered. Some, however, must be sown 
at once, or they will not germinate at all. The spores of the 
common Osmunda regalis, and probably of the other members of 
the genus, must be sown as soon as ripe, or they fail to germinate. 
The prothallia of Osmunda regalis, if carefully covered with 
glass, may be kept for a long time. Prothallia of this fern in 
the writer's laboratory produced ribbon-like outgrowths three- 
sixteenths of an inch wide, and often more than two inches in 
length. These prothallia continued to produce archegonia, 
antheridia, and the ribbon-like outgrowths for more than a year, 

103 



104 Methods in Plant Histology 

when they suddenly "damped off." Pteris aquilina and many 
other ferns often furnish a good supply of antheridia in three 
weeks after sowing, and the archegonia appear soon after, but it 
is well to make sowings six weeks before material is needed for 
use. In Pteris aquilina and in many others, if the spores are 
sown too thickly, only antheridial plants will be obtained. If 
prothallia are to produce archegonia, they must have sufficient 
room and nutrition. If there are no greenhouse facilities and 
the prothallia must be grown in the laboratory, it is a good plan 
to take a glass dish, ten or twelve inches in diameter and about 
two inches deep, put a layer of broken pieces of flower pots on 
the bottom, cover this with a layer of rich loam, and over this 
sprinkle a layer of fine, clean sand, since sand is much more 
easily washed away from the rhizoids than is the loam. The 
whole should now be thoroughly wet, but not so as to have 
water standing on the bottom. Sow the spores and cover with 
a tightly fitting pane of ground glass. There should be no 
need for moistening the culture again, for prothallia can be kept 
fresh and vigorous for several months, or even for a year, without 
any wetting. When it is desired to secure fertilized material, 
sprinkle the prothallia with water, and the young sporophytes 
will soon appear. If greenhouse facilities are available, any gar- 
dener can grow prothallia in abundance without any directions 
from those who want the material. 

The peculiar tuberous prothallia of Botrychium are seldom 
found except by the experienced collector. The older prothal- 
lia, however, may be found by anyone who is able to recognize 
Botrychium when he sees it. Dig up young plants not more 
than three or four inches in height, and the prothallia, which 
persist for years, will often be found still attached. They are 
easy to cut and may be handled like other prothallia. 

Fern prothallia of the usual type are excellent objects for 
testing fixing agents, since the prothallia, while still in the fixing 
agent, may be examined with the microscope, and fluids which 
cause plasmolysis may be rejected. It will sometimes happen 
that plasmolysis may be avoided by varying the proportions of 



Pteridophytes 



05 



the ingredients of a fixing agent. Chromo-acetic acid with 
about 0.6 g. chromic acid and 0.4 cc. of acetic acid to 100 cc. of 
water will seldom cause plasmolysis, and will usually insure good 
fixing. It is a mistake to suppose that because prothallia are 
suchdelicateobjects 
the fixing will take 
but a few minutes. 
We should recom- 
mend at least twen- 
ty-four hours in 
chromo-acetic or 
Flemming's fluid 
and two or three 
days will do no harm 
and may be better. 
If hot corrosive sub- 
limate-acetic acid or 
hot picro-acetic be 
used, the fixing re- 
quires only two or 
three minutes, but 
results are not as 
uniformly success- 
ful as with members 
of the chromic-acid 
series. After any 
of the chromic-acid 
series, two or three 
hours' washing in 
water will be suffi- 
cient, if the water be changed as often as it becomes in the 
least degree discolored. 

If preparations are to be mounted whole, as shown in fig. 52, 
they should be stained as soon as the washing is finished. Any 
of the following methods gives good results : 

a. Stain in Mayer's haem-alum six hours or over night, 




Fig. 52. Pteris aquilina. 
A , filamentous stage. B, the apical cell has been established and 
several segments have been cut off. The figure shows the initial 
rhizoid, and also three rhizoids coming from the main body of the 
prothallium. C, an older prothallium covered with antheridia in vari- 
ous stages of development. From a glycerine mount, fixed in chromo- 
acetic acid and stained in Delafield's hsematoxylin. (Miss M. E. 
Tarrant.) 



io6 



Methods in Plant Histology 



wash in water one or two hours, and transfer to 10 per cent, 
glycerine. 

b. Stain in Delafield's hematoxylin thirty minutes, wash in 
water one or two hours, decolorize in water acidulated with 
hydrochloric acid (3 drops of HC1 to 100 cc. water) one to 




Fig. 53. Pteris cretica. X 250. 



A, early stage in the development of the archegonium. B, later stage showing the oosphere, ventral 
canal cell, and three nuclei in the neck canal. C, still later stage almost ready for fertilization. The 
ventral and neck canal cells are breaking down, and the oosphere is nearly mature. Cells surrounding the 
oosphere have become richer in protoplasmic contents, and stain more deeply. D, first division of the 
embryo. E, young embryo still showing the outlines of the four quadrants. The apical cell in the lower 
left quadrant has cut off the first layer of the root cap. All drawn from material stained in bulk in alum 
carmine, a method not to be recommended. 

thirty minutes — the time can be determined only by experi- 
ment — wash in water until the rich purple color of the haema- 
toxylin replaces the red due to the acid, and then place in 10 per 
cent, glycerine. 

c. Stain with Delafield's hematoxylin as in b, and after the 
last washing in water stain two to four minutes with an aqueous 





Pteridophytes 107 

solution of eosin, wash thoroughly in water, and transfer to the 
10 per cent, glycerine. Instead of using the acid alcohol after 
the haematoxylin, the eosin may be allowed to act for several 
hours, and then 1 per cent, acetic acid may be used for a few 
minutes. Wash very thoroughly in water, and, if the stains 
appear satisfactory, transfer to 10 per cent, glycerine as before. 

d. Use a 2 per cent, solution of iron alum two hours, wash 
in water five minutes, stain in y 2 per cent, haematoxylin two to 
six hours, wash in water five minutes, and then treat again with 
iron alum until the stain is sat- 
isfactory. Wash thoroughly in 
water and transfer to 10 per 
cent, glycerine, which, as usual,, 
will concentrate sufficiently for 
mounting in three or four days. 

For paraffin sections such A 

1 • n 1 Fig. 54. Pteris cretica. 

as are shown in figs. 5? and 54, . , . 

o ~>^/ ~> 1 a , section of a nearly mature antheridium showing 

the material Should be paSSed the anthenwoids inside X 333. B, a developing 

1 antherozoid showing the blepharoplast drawn out 

thrOllCrh the alrohnls allnwincr into a dee P lv staining band. X 1900. Fixed in 
UirUUgll U1C dlCOllUlS, dllOWUlg chromo . acetic acid and stamed in Haidenhain's iron 

about three or four hours for ^ m - h ™° x y lin - 

each grade. The mixtures of xylol and absolute alcohol should 
take about six hours, and as soon as the pure xylol has been 
added, the piece of paraffin may be added at the same time. In 
the bath two or three hours will give good results. It would be 
worth while to determine the duration of the bath for such objects. 
Some workers claim that ten or fifteen minutes is amply suffi- 
cient, and that there is less danger from shrinking, while others 
think that several hours is better, and that two or three days will 
do no damage, if the fixing has been thorough and the tempera- 
ture is not allowed to become higher than 50 C. If bergamot 
oil be used instead of xylol, there is less danger of collapse pre- 
vious to the bath, but the bath itself must usually be more pro- 
longed, since the bergamot oil is not so easily gotten rid of as is 
the xylol. 

For morphological purposes sections 15 /jl to 20 ^ thick are 
better than thinner ones. Delafield's haematoxylin, with or without 



io8 



Methods i?i Plant Histology 




a counter-stain with erythrosin, is good for such sections as 
are represented in fig. 5j. If very thin sections are wanted for 
cytological study, it is better to use the safranin-gentian violet- 
orange combination. For such views of antheridia as are shown 
in fig. j4, Haidenhain's iron alum-haematoxylin seems to bring 
out the blepharoplast (centrosome) most sharply. 

The Sporophyte. — Methods for young sporophytes like those 
shown in fig. jj, D and E, are the same as for archegonia and 
antheridia. 

Roottips to show the prominent apical cell are easily imbedded 
in paraffin. They should be cut 15 \x to 20 /u, thick. Delafield's 

hematoxylin, without any con- 
trast stain, is best for bringing 
out the prominent apical cell and 
its segments. 

Sporangia should also be cut 
in paraffin. Pteris is a good form 
for sporangia, since the long mar- 
ginal sorus makes it possible to 
get an immense number of median 
longitudinal sections of sporangia 
in a single preparation. (Fig. 55.) 
Pteris ere tic a can always be 
found in fruit in greenhouses. 
Select a series of stages. The leaf should be cut with a razor — 
not with scissors — into pieces about one-fourth of an inch long. 
If sections of the whole leaf are not wanted, only the marginal 
sorus need be cut off; in this way a much greater number of 
sporangia may be gotten upon a slide. Aspidium and Cyrtomium 
give beautiful views of the indusium covering the cluster of 
sporangia. The most satisfactory preparations will be gotten 
from material in which the sporangia have not yet begun to 
turn brown. 

Botrychium furnishes excellent and usually accessible material 
for studying the development of sporangia of the eusporangiate 
type. For the archesporium and early stages in the development 



A 




Fig. 55. Pteris cretica. X 560. 

A, young stage in the development of the spo- 
rangium. B, older stage showing the tapetum. 
Fixed in chromo- acetic acid and stained in 
Delafield's hsematoxylin and erythrosin. 



Pteridophytes 109 

of the sporangia the material should be collected in the latter 
part of August or in September. When the sporangia for one 
year are ready to shed their spores, the sporangia for the next 
year will be found in early stages of development. When a 
frond is found, dig the plant up very carefully and remove the 
large frond. At its base will be found the frond for the next 
season, the sterile portion bent over so that its tip is directed 
downward and the fertile portion — one-fourth to three-fourths 
of an inch in length — projecting from the ventral surface of the 
sterile frond and directed upward. It is worth while to preserve 
the whole bud and also portions of the stipe, rhizome, and roots. 
These vegetative parts should be cut, with a razor or very sharp 
knife, into pieces about one-fourth of an inch in length. The 
larger roots are better for transverse sections. The root tips are 
unusually favorable for preparations of the apical cell. The 
upper part of the rhizome cuts easily, but all parts of this plant, 
even the older portions of the rhizome, can be cut in paraffin. 
Delafield's haematoxylin or iron alum-hsematoxylin gives fine 
preparations of the young sporangia. The older sporangia, from 
the mother-cell stage up to the shedding of the spores, stain 
better in cyanin and erythrosin or in the safranin-gentian violet- 
orange combination. The following schedule for paraffin sections 
will give elegant mounts of the root, stipe, and rhizome : 

1. Stain in safranin, about 24 hours. 

2. Fifty per cent, alcohol until little or no safranin is left in the cellulose 
walls. Use a very weak acid alcohol, if necessary. 

3. Delafield's haematoxylin, 5 to 10 minutes. 
\. Water, 15 minutes. 

5. Thirty-five and 50 per cent, alcohol, 10 seconds each. 

6. Acid alcohol — the same as was used for the safranin — a few seconds. 

7. Seventy per cent, alcohol until the purple color returns. The lignified 
walls should now show a brilliant red and the cellulose walls a rich purple. 
If either stain is too deep or too faint, do not proceed any farther before 
making the necessary correction. If the safranin washes out, and the 
haematoxylin is too intense, do not use any acid alcohol in the second 
step and shorten the period in the haematoxylin. 

8. Eighty-five and 95 per cent, alcohol, a few seconds each. 

9. Hundred per cent, alcohol, 30 seconds to 1 minute. 

10. Xylol until cleared. 

11. Balsam. 



no 



Methods in Plant Histology 



The spore mother-cells of Osmunda are excellent for a study 
of mitosis. The young sporangia of 0. cinnamomea and 0. Clayto- 
niana show the mother-cell stage in the autumn, but the division 
into spores does not occur until the following spring, in the 
vicinity of Chicago the mitotic figures being found during the 

latter part of April. 0. 
regalis does not reach the 
mother-cell stage in the 
autumn. Material for 
mitosis should be col- 
lected during the first 
two weeks in May. The 
material may be fixed in 
the medium chromo- 
aceticsolutionorinFlem- 
ming's weaker solution. 
Sections should not be 
thicker than io /x, and 
5 ix will be found more 
satisfactory. 

Preparations of the 
woody structure of the 
sporophyte are easily 
made. The rhizome of 
Pteris aquilina affords as 
good material as any 

Fig. 56. Pteris aquilina. {fig'5 6 )- In digging Up 

A part of a transverse section of the vascular bundle of the rhizomes, do not merely 

rhizome. X 166. e, endodermis. /, pericycle. st, sieve tube. ^ 

t, seal ari form tracheid. Drawn from a celloidin section from dlSf down Until the rhi- 

material fixed in picro-acetic acid and stained in safranin and ° 

Delafield's hematoxylin. zome can bg g raS p e d and 

then pull it up, for such material is sure to show the pericycle of the 
bundles torn away from the parenchyma. Dig carefully around 
the rhizome and then cut off with a very sharp knife pieces about 
two inches in length. Put the fresh rhizome into the hand micro- 
tome and cut as thin sections as possible. Keep the knife wet 
with 95 per cent, alcohol and put the sections into 95 per cent. 




Pteridophytes 



1 1 1 




alcohol as fast as they are cut. Fifteen or twenty minutes is 

sufficient for fixing. Pass down through the grades of alcohol, 

about two or three minutes in each grade. Stain in safranin 

twenty-four hours, wash in water ten minutes, and then stain in 

Delafield's hematoxylin ten to twenty minutes. Pass through 

the alcohols, about one minute in each grade, at 70 per cent. 

alcohol transferring to acid alcohol from one to five seconds ; 

then pass on, clear in 

xylol, and mount in 

balsam. If the stain 

is successful, the 

xylem should show a 

brilliant red, and the 

cellulose walls a rich 

purple. 

Another method is 
to stain for about twen- lX\ t/T 'J^%s%Jd( r 
ty minutes, or several .~w:^^. 

, .... Fig. 57. Marsilea quadrifolia. X 333. 

hours, in iodine green . . .,.,.. • ■ *. • -\ 

A , apex of megaspore with archegonium containing the oosphere. 

r*r mp>tViA7l crrf*f±r\ rincf* Large starch grains are shown beneath the archegonium. B, young 
UI illCLliyi gXCCll, IlilbC embry0 _ Both fixed in picro- acetic acid and stained in Delafield's 

in 70 per cent, alcohol hjEmatox y lin - < Mlss M - E - tarkant.) 
until the green is prominent only in the xylem, then stain for 
from one to three minutes in acid fuchsin (i per cent, solution 
in 70 per cent, alcohol), dehydrate rapidly, clear in xylol, and 
mount in balsam. The xylem should have a sharp bright green 
color, and the cellulose a bright red. 

The most beautiful preparations may be obtained by imbed- 
ding in celloidin and staining in safranin and Delafield's hema- 
toxylin. These stains allow a use of acid which extracts all color 
from the celloidin and still leaves a sufficient amount in the 
tissues. The large apical cells of rhizomes are easily cut in 
either paraffin or celloidin. 

The megaspores and microspores of Marsilea are easily 
obtained [figs. 57 and 5c?). Cut away a portion of the hard 
sporocarp and place the sporocarp in a dish of water. The 
gelatinous ring with its sori will sometimes come out in a few 



12 



Methods in Pla?it Histology 





minutes. In less than twenty-four hours the microspores, start- 
ing from the one-cell stage, will produce the mature antherozoids. 
The development of the megaspore is equally rapid. Embryos 
are abundant in two or three days. For morphological work, 
picro-acetic acid, used hot, is very good, since the material does 
not occasion so much difficulty in cutting. Chromo-acetic 
material allows better staining, but the cutting is more uncertain. 

It is best to prick the 
megaspores with a 
needle while they are 
in the fixing fluid, in 
order to facilitate the 
infiltration of paraffin. 
Better mounts of the 
microspores can be 
obtained if the trou- 
blesome megaspores 
be picked out from 
the sorus while the 
material is still in the fixing agent or the alcohols. The mega- 
spores must be imbedded in rather hard paraffin, and one must 
expect to hone the knife thoroughly before it can be used again, 
for when the knife strikes a megaspore of one of the heterospor- 
ous pteridophytes, it seems like striking a grain of sand. 

The following schedule will usually give good sections of the 
hard megaspores of the heterosporous pteridophytes, whether 
they are to be cut separately or in their sori or strobili. The 
essential features of the method are suggested by Miss F. M. 
Lyon's work on Selaginella: 

i. Chromo-acetic acid, medium solution, 2 to 6 days. 

2. Wash in water, 1 day. 

3. Thirty-five to 95 per cent, alcohol, 1 day each. 

4. One hundred per cent, alcohol, 3 or 4 days, changing several times. 

5. Mixtures of alcohol and xylol, 2 days. 

6. Pure xylol at 53 C, 1 or 2 days. 

7. Add paraffin to the xylol and keep at 53 ° C. for 2 or 3 days. 



Fig. 58. Marsilea quadrifolia. X 560. 



A , microspore before germination. B, microspore with antheridia 
nearly mature. Fixed in chromo-acetic acid and stained in safranin- 
gentian violet-orange. 



Pteridophytes 1 1 3 

8. Pure paraffin 53 C, 3 or 4 days, and then in harder paraffin at 6o° C. to 
70 C, 2 or 3 days. 

9. Imbed in rather thin cakes. 

While the method is tedious, it is worth the trouble, for even 
old strobili of Selagi?iella yield smooth ribbons at 5 ^. 

As Jig. 57 suggests, the mature archegonia, and especially the 
young embryos, may be removed from the top of the megaspore 
and cut with perfect ease. 

The spermatozoid, which in Marsilea has an unusually large 
number of turns in the spiral, may be mounted by methods 
already described. 



CHAPTER XVI. 

PTERIDOPHYTES. 

EQUISITINE^. 

The prothallia of Equisetum are easily grown by the method 
already described for the Filicinese, but the spores must be 
sown as soon as ripe, because they fail to germinate if kept more 
than a few days. In the vicinity of Chicago the spores of 
Equisetum arvense are shed during the latter part of April. The 
methods for preparing the prothallia are the same as for the 
Filicinese. The sporangia are harder to cut, but good prepara- 
tions should be secured from paraffin material. E. arvense is 
abundant everywhere, and is to be preferred on account of the 
comparative ease with which the sporangia and other portions of 
the fertile shoot can be cut. Longitudinal sections of the 
younger strobili show various stages in the development of the 
spores, the more advanced stages being found at the base of the 
strobilus. Tetrads may be found at the base of the strobilus, 
while the spore mother-cells at the apex are still undivided. Of 
course, it is impossible to stain a longitudinal section of such a 
strobilus so that all stages will be satisfactory. For the beginner, 
at least, this is not a serious objection, for he will be almost sure 
to secure some stage beautifully stained. The experienced 
worker, who is able to control his staining with more precision, 
will prefer transverse sections. Material for sporangia should 
be obtained as soon as the fertile shoot appears above ground, 
and if it can be obtained earlier, so much the better. When the 
spores are shed, the young sporangia which are to develop the next 
year can already be detected. The safranin-gentian violet-orange 
combination can be recommended for the development of the 
mother-cell and the formation of tetrads, but Delafield's hema- 
toxylin or Haidenhain's iron alum-haematoxylin will be more 
satisfactory for earlier stages. 

115 



1 1 6 Methods i?i Plant Histology 

The roots are very small, but have large cells and easily yield 
good preparations. In case of such small objects it is a good 
plan to add a few drops of eosin to the alcohol during the pro- 
cess of dehydrating, in order that the material may be seen more 
easily. The slight staining does no damage, even if more 
critical stains are to be used after the sections are cut. It is 
easy to get longitudinal sections of the roots by cutting trans- 
verse sections of the nodes. In E. arvense these roots at the nodes 
are quite numerous. Sections of the stem of the fertile shoot of 
E. arvense are easily cut in paraffin or celloidin, but sections of 
the stem of E. hiemale or similar species do not cut in paraffin, 
and results are rather uncertain even in celloidin. The growing 
points of stems, however, may be cut with ease in paraffin. E. 
arvense is particularly favorable on account of the numerous 
apical cells which may be found in a single preparation. Dela- 
field's hematoxylin, used alone, is good for the apical cells, but 
for sections of older stems a slight counter-stain with erythrosin 
will improve the mount. 



CHAPTER XVII 



PTERIDOPHYTES. 
LYCOPODINE^. 

It is not very difficult to get paraffin sections of young 
sporangia of Selaginella, but the method just recommended for 
Marsilea should be resorted to for the older strobili {fig. 59). 

The older megaspores had better be pricked with a needle 
and cut one at a time. A slight puncture at the basal portion of 
the megaspore 
does no damage 
and insures a thor- 
ough infiltration. 
If the megaspores 
are imbedded sep- 
arately, they will 
usually orient 
themselves so that 
sections perpen- 
dicular to the par- 
affin cake will 
show the most in- 
structive views of 
the gametophyte 
structures. 

Delafield's has- 

matoxylin, used alone, is a good stain. The safranin-gentian 
violet-orange combination gives a brilliant differentiation in 
stages like fig. 59, B. The microspores of Isoetes offer the same 
difficulties. Cyanin and erythrosin is a fine combination for the 
reserve food-stuffs in Isoetes macrospores. 

For young sporangia of Isoetes, the leaves should be cut off 
about one-eighth of an inch above the sporangia, and the stem 

117 




Fig. 59. Selaginella Mertensii. X 93. 

A, microsporangium containing microspores. B, a megaspore show- 
ing the beginning of the prothallium. Fixed in picro-acetic acid and 
stained in Delafield's haematoxylin. (Miss M. E. Tarrant.) 



1 1 8 Methods in Pla?it Histology 

should be cut off below, leaving just enough to hold the 
sporangia together. Cut longitudinal sections and stain in 
Delafield's hematoxylin, with or without the addition of a light 
touch of erythrosin. The older sporangia had better be removed 
and cut separately. Transverse sections of the stem are very 
interesting. Young stems and the apices of old ones cut well 
in paraffin, but older stems will give more satisfactory results in 
celloidin. 

It is much easier to get good preparations of the sporangia 
of Lycopodium, since there are no megaspores with their hard 
walls. Delafield's hematoxylin, without any contrast stain, will 
bring out the developing sporangia, which are usually to be 
found even after the sporangia in the lower part of the strobilus 
are beginning to shed their spores. The directions for the 
rhizome of the Filicineae will also serve for the stem of 
Lye op odium. 



CHAPTER XVIII. 

SPERMATOPHYTES. 

In this immense group we cannot hope to give even approxi- 
mately complete directions for making preparations, but must 
be content to give a few hints which may prove helpful in col- 
lecting material and in securing mounts of the more important 
structures of the flowering plants. We shall consider the gym- 
nosperms and the angiosperms separately, although in many 
respects the technique is the same for both. 

GYMNOSPERMS. 

Since Pinus is a characteristic type, we shall describe methods 
for demonstrating various phases in the life-history of this genus, 
hoping that the directions will enable the student to experi- 
ment intelligently with other forms. 

Spermatogenesis. — In October the clusters of staminate cones 
which are to shed their pollen in the coming spring are already 
quite conspicuous. The cones should be picked off separately, 
and the scales should be carefully removed so as to expose the 
delicate greenish cone within. At this time the archesporial 
cells are easily distinguished. Material collected in January, or 
at any time before growth is resumed in the spring, shows about 
the same stage of development. If it is desired to secure a 
series of stages with the least possible delay, a branch bearing 
numerous clusters of cones may be brought into the laboratory 
and placed in a jar of water. Growth is more satisfactory in 
case of branches broken off in the winter than in those brought 
in before there has been any period of rest. The material can 
be examined from time to time, and a complete series is easily 
secured. The karyokinetic figures in the pollen mother-cells 
furnish exceptionally instructive preparations. Staminate cones 
which will yield karyokinetic figures can be selected with con- 
siderable certainty by examining the fresh material. Crush a 

119 



120 Methods in Plant Histology 

microsporangium from the top of the cone and one from the 
bottom, add a small drop of water and a cover to each, and exam- 
ine. If there are pollen tetrads at the bottom, but only undi- 
vided spore mother-cells at the top, it is very probable that 
longitudinal sections of the cone will yield the figures. If a drop 
of methyl green be allowed to run under the cover, it will enable 
one to see whether figures are present or not. When desirable 
cones are found, they should be cut longitudinally into two 
halves. The later stages, showing the germination of the micro- 
spores, furnish better sections if the cones are cut transversely 
into small pieces about three-sixteenths of an inch thick. It is 
very easy to get excellent mounts of the pollen just at the time of 
shedding. Shake a large number of cones over a piece of paper, 
thus securing an abundance of material. Fix in chromo-acetic 
acid, wash in water (a few minutes is sufficient, and the water need 
not be changed), pass through the alcohols, allowing each to 
act for about two hours, make the usual gradual transition from 
alcohol to xylol, and from xylol to paraffin. It is best that the 
material should be in a small bottle not more than one-fourth 
of an inch in diameter ; at any rate, the pollen should be in such 
a bottle during infiltration, which should not require more than 
two or three hours, although a longer period does no harm if the 
temperature does not rise above 52 or 53 C. Now put the lower 
portion of the bottle into cold water, and thus harden the paraffin 
as quickly as possible. Break the bottle carefully, cut off the lower 
portion of the paraffin containing the pollen, mount it on a block 
in the usual manner, and trim away some of the paraffin so that 
two parallel surfaces will make the sections ribbon well. Material 
in this stage shows a large tube nucleus, a somewhat lenticular 
cell with a more deeply staining nucleus, and, lastly, two small 
prothallial cells quite close to the spore wall. The prothallial 
cells cannot always be detected at this stage, and there may be 
some doubt as to whether two such cells are always present. 
The division of the lenticular cell into "stalk cell" and "body 
cell," and also the division of the body cell into the two male 
cells, must be looked for in sections of the nucellus of the ovule. 



Spermatophytes 



121 



Oogenesis. — The entire ovulate cone at the time of pollina- 
tion is easily cut in paraffin. Longitudinal sections of the cone 
at this time give good views of the bract and ovuliferous scale 
bearing the ovules. The integument is very well marked, and 
in the nucellus one or more sporogenous cells can usually be 
distinguished. As soon as the scales close up after pollination, 
the cone will be too hard to cut, and it will be necessary to 
remove the scales and cut them separately. For a study of the 
ovule and the structures within it, better preparations will be 





A b 



JOWLX fVpp. H»«" 





Fig. 60. Pinus Laricio. X 104. 



A, top of prothallium with an archegonium just before the cutting off of the ventral canal cell. 
Fixed in Flemming's weaker solution and stained with Haidenhain's iron alum-hsematoxylin. Collected 
June 18, 1897. B, C, and D, early stages in the formation of the embryo. Fixed in chromo-acetic acid, 
and stained in safranin -gentian violet-orange. Collected July 2, 1897. 

obtained by carefully cutting off the pair of ovules from the 
scale. For preparations like that represented in. fig. 60, A, it is 
a good plan to remove the endosperm with its archegonia from 
the ovule. Fixing, infiltration, and cutting will then occasion 
but little trouble, and the whole ribbon may be gotten upon a 
single slide. However, at this stage the pollen tubes with their 
contents are rapidly working their way through the nucellus 
toward the archegonia, and consequently it is better to retain 



122 Methods in Plant Histology 

enough of the tissues of the ovule to keep the nucellus in place. 
In later stages, after fertilization has taken place, it is necessary 
to remove the endosperm. In stages like fig. 60, B, C, D, and 
later, the developing testa should be dissected away with great 
care, for a very slight pressure is sufficient to injure the delicate 
parts within. Mature embryos may be dissected out from the 
endosperm before fixing, but it is hardly necessary, since they 
cut quite well if left in place. The "pine nuts" or "pifion," to 
be found upon the market, are good for a study of the mature 
embryo. The testa, which is quite a hard shell, should be taken 
off, and the endosperm should be allowed to soak in water for 
about twenty-four hours, after which the embryo may be dis- 
sected out and fixed. 

The period at which the various stages may be found varies 
with the species, the locality, and the season. In Pinus Laricio 
(the common Austrian pine) at Chicago, in the season of 1897, 
material collected May 27 did not yet show archegonia ; the 
ventral canal cell was cut off about June 21 (see fig. 61), the 
fusion of the pronuclei occurred about a week later, and stages 
like fig. 60, B, C, and D, were common in material collected July 
2. In the season of 1896 all the stages appeared about two 
weeks earlier. In Pinus sylvestris the stages appeared a little 
earlier than in Pinus Laricio. After the stage shown in fig. 60, A, 
has appeared, it is necessary to collect at intervals of not more 
than two days until the stage shown in fig. 60, D, is reached. If 
collections are made at intervals of four or five days, the most 
interesting stages, like the cutting off of the ventral canal cell, 
fertilization, and the first divisions of the nucleus of the oospore, 
may be missed altogether. It should be mentioned that all the 
ovules of a cone will be in very nearly the same stage of devel- 
opment. 

A rather strong chromo-acetic acid (1 g. chromic acid and 
y 2 cc. glacial acetic acid to 100 cc. water) can be recom- 
mended for the entire series in spermatogenesis, oogenesis, fer- 
tilization, and formation of the embryo. After repeated trials 
the popular Flemming's solution does not seem to be at all 



Spermatophytes 



23 



superior and often fails to give as good results as the cheaper 
fixing agent. 

The following stains may be suggested : for studying the 
pollen tubes in the nucellus, cyanin and erythrosin ; for the 
development of the archegonium up to the stage shown in 
fig. 60, A, 
Delaf ield's 
hematoxy- 
lin; for the 
stages shown 
in fig. 60, A, 
iron alum- 
haematoxylin 
or the safra- 
nin - gentian 
violet-orange 
combination; 
for the stage 
shown in fig. 
6i, nothing 
seems to 
equal the 
safranin-gen- 
tian violet- 
orange com- 
bination ; for 
stages like 
fig. 60, B, C, 
and D, and 
also for later 
stages in the 
development 
of the em- 
bryo, Delafield's hematoxylin brings out the walls perfectly, but 
since mitotic figures are very frequent in these stages, it is worth 
while to use the safranin combination with some preparations, 




Fig. 61. Pinus Laricio. X 710. 

The mitotic figure concerned in cutting off the ventral canal cell. The 
nucleus at the lower end of the spindle is the nucleus of the ob'sphore. 
Fixed in chromo-acetic acid, and stained in the safranin-gentian violet- 
orange combination. Collected June 21, 1897. 



124 Methods in Plant Histology 

although it is much inferior to Delafield's haematoxylin when 
cellulose walls are to be emphasized. 

All the stages which have been described can be cut in 
paraffin with little difficulty. 

The Leaves. — The leaves of our common gymnosperms cut 
readily in paraffin while they are young and tender, but as they 
approach maturity it is a fruitless task to attempt paraffin sec- 
tions. Celloidin sections are far more satisfactory. Cut the 
needles into pieces about one-fourth of an inch long, fix in a 
picro- corrosive -acetic mixture ( y 2 g. picric acid, 2 g. cor- 
rosive sublimate, I cc. glacial acetic acid, ioo cc. 50 per 
cent, alcohol). If used hot, five minutes is sufficient, but if 
used cold it should be allowed to act for two or three hours. 
After the material has been imbedded in celloidin, the block 
should be placed in equal parts of 95 per cent, alcohol and 
glycerine for a few days, after which it should cut quite readily. 
Stain with safranin and Delafield's haematoxylin, clear in Eycle- 
shymer's clearing mixture, and mount in balsam. 

Fairly good sections may be obtained in great quantities with 
little trouble by the following method : Make a bunch of the 
needles as large as one's little finger, wrap them firmly together 
with a string, allowing about an eighth of an inch of the bunch 
to project above the wrapping ; then fasten the whole in a hand 
microtome, and every stroke of the razor will give twenty or 
thirty sections, some of which will surely be good. As the sec- 
tions are cut, they maybe put directly into 95 per cent, alcohol, 
and after a few minutes can be transferred to 50 per cent, alco- 
hol and then to the stain. Dehydrate, clear in xylol, and mount 
in balsam. 

Stems and Roots. — With a sharp razor fairly good sections 
of stems and roots may be made without imbedding, especially 
if the hand microtome be used. Young buds may be cut in 
paraffin. Stems and roots as large as half an inch in diameter 
can be cut in celloidin. The material should be cut into pieces 
not more than one-fourth of an inch long. The following treat- 
ment should give good results : 



Spermatophytes 1 2 5 

1. Picro-corrosive-acetic mixture, five minutes if used hot, or 
wo to three hours if used cold. 

2. Wash in 50 per cent, alcohol, to which a little iodine has 
been added, two hours; 70 per cent., 8.5 per cent., 95 per cent., 

our hours each ; absolute alcohol, ten hours ; then change to 
fresh absolute alcohol, which should act for ten hours longer. 

3. Ether alcohol, twenty-four hours. Some prefer to pre- 
cede the ether alcohol by a mixture of equal parts of ether 
alcohol and absolute alcohol. 

4. Thin celloidin (about 2 per cent.), two or three days; 
6 per cent, celloidin, two or three days; 10 per cent, celloidin, 
two or three days. After the thin celloidin has acted for a few 
days, the cork may be removed for a short time each day, thus 
allowing the thin celloidin to become thick by the evaporation 
of the ether alcohol. 

5. Get some small blocks of wood (three-eighths inch cubes 
of white pine are good), wet one of them in ether alcohol, dip it 
into thin celloidin, place the object upon the block in convenient 
position for cutting, pour over it a few drops of 10 per cent, cel- 
loidin, and then plunge the whole into chloroform. Leave it in 
the chloroform about twenty-four hours, and then transfer to a 
mixture of equal parts of 95 per cent, alcohol and glycerine, 
where it should remain for several days. Material may be kept 
here indefinitely. Even refractory stems may be cut after they 
have been in this mixture for a couple of weeks. 

6. Cut the sections, keeping the knife wet with the alcohol 
and glycerine mixture. Transfer the sections to 70 per cent, 
alcohol, then to 50 per cent., and then to the stain. 

7. Stain in safranin, twenty-four hours; wash in 35 per cent, 
alcohol, about a minute; stain in Delafield's hematoxylin, five to 
ten minutes; wash in water, two minutes ; 35 per cent, alcohol, two 
to five minutes ; 50 per cent, alcohol, two to five minutes ; acid 
alcohol, one to ten seconds; 70 per cent., 85 per cent., 95 per 
cent., about two minutes each ; Eycleshymer's clearing mixture 
until cleared, usually about one or two minutes. Mount in 
balsam. 



126 



Methods in Plant Histology 






■JVUnVoersity of Qjieoflo. 



ioT^.rruL.loiivj.ttti*}.. 



&to. 



TUcutsut . 



^ 



■■>■::. -ZMMMMMr 



JOOLX. F\pp. A\v<!-. 



Xylem should show a brilliant red color and cellulose a 
rich purple, if the stain is successful. If either stain is too weak 
or too prominent, the duration of the stain, the length of time 
in the alcohols, or the time in acid alcohol must be varied until 
the desired result is secured. 

People who make all their anatomical sections without imbed- 
ding may regard this method as tedious and unnecessary, but 

such preparations 
will show much 
which is neverseen 
in mere free-hand 
sections, for the 
reason that free- 
hand sections, if 
thin enough to 
show any detail, 
will lose most of 
their cell contents, 
while in celloidin 
sections every- 
thing is held in 
place. Even if the 
celloidin sections 
be passed through absolute alcohol and cleared with clove oil, a 
process which dissolves away the celloidin, the contents of the 
cells will still be retained in most cases, and stains which 
cannot be extracted from celloidin may be used. However, 
we prefer to use the safranin and Delafield's hematoxylin and 
retain the celloidin, since this combination can be extracted 
completely from the celloidin and still leave the object brilliantly 
stained. 

For preparations of the mature wood a piece of white pine 
from a dry-goods box furnishes perfect material. It should not 
be imbedded, but the cutting will be facilitated if the piece be 
soaked for a few hours in water or in the alcohol and glycerine 
mixture. In all preparations designed to show the structure of 



THE 

lv*c{ - jt&wq. ttcA/. 



"MJe 



1 



@ a 

I 1 
I 1 



Fig. 62. 
Slides showing labels and methods of arranging sections. 



Spermatophytes 127 

wood there should be three sections — a transverse, a longitu- 
dinal radial, and a longitudinal tangential. These may be 
arranged as in the upper slide of fig. 62, but if it is desired to 
make a thorough study of the structure, it is a good plan to have 
on each slide several sections of each kind, thus having an 
opportunity to use a variety of stains. Fuchsin and iodine green 
is a good combination. The safranin and Delafield's haema- 
toxylin is also excellent. 

For a study of young stems or roots, preparations like that 
shown in the lower slide of fig. 62 will be found very convenient. 



CHAPTER XIX. 

SPERMATOPHYTES. 
ANGIOSPERMS. 

Success depends largely upon judgment and care in selecting 
and trimming material before it is put into the fixing agent in 
the field. While the following directions cannot be applied to 
all plants, they should, nevertheless, enable the student to make 
such modifications as may be demanded by any particular form 

Floral Development. — For a study of floral development very 
young buds are necessary, and it is best to select those forms 
which have rather dense clusters of flowers, in order that a com- 
plete series may be obtained with as little trouble as possible. 

The usual order of appearance of floral parts is (i) calyx, 
(2) corolla, (3) stamens, and (4} carpels, but if any of these 
organs are reduced or metamorphosed, their order of appearance 
may be affected. 

Floral development is easily studied in the common Capsella 
bursa-pastoris. The best time to collect material is late in March 
or early in April. Dig up the plant, carefully remove the leaves, 
and in the center of the rosette a tiny white axis will be found. 
A series of these axes from one-eighth of an inch to three- 
eighths of an inch in length and from one-sixteenth of an inch 
to three-sixteenths of an inch in diameter will give a very com- 
plete series of stages in the development of the floral organs. 
Preparations from the apex of the shoot taken after the inflores- 
cence appears above ground are not to be compared with these 
taken early in the season. Fix in chromo-acetic acid and stain 
in Delafields hsematoxylin. The sections should be longitudinal 
and about 5 \i thick. 

The common dandelion, Taraxacum officinale, affords an excel- 
lent series with little labor. Examine vigorous plants which have, 
as yet, no flowers or buds in sight. Dig up the plant and dissect 
away the leaves. If there is a white cluster of flower buds, the 

129 



130 Methods i?i Plant Histology 

largest not more than three-sixteenths of an inch in diameter, 
cut out the cluster, leaving only enough tissue at the base to 
hold the buds in place. Larger heads should be cut separately. 
Fix and stain as in Capsella. 

Our most common thistle, Cnicus lanceolatus, shows the floral 
development with unusual clearness, but the preparation of the 
material is somewhat tedious. The involucre, which is too hard 
to cut, must be carefully dissected away. Retain only enough of 
the receptacle to hold the developing florets in place. A series 
of sizes with discs varying from one-eighth of an inch to three- 
eighths of an inch in diameter will show the development from 
the undifferentiated papilla up to the appearance of the arche- 
sporial cell in the nucellus of the ovule. The Canada thistle, 
C?iicus arvensis, is equally good, but it is more difficult to dissect 
out the desirable parts. 

In the willows, Salix, the bud scales must be removed and 
the copious hairs should be trimmed off as much as possible 
with scissors, after which the catkin should be cut in two longi- 
tudinally and placed in the fixing agent. 

Spermatogenesis. — The earlier stages in spermatogenesis will 
be found in the preparations of floral development. For tracing 
the nuclear changes involved in this process the lilies furnish 
very good material, because the cells and nuclei are exceptionally 
large. Several species of Lilium are common in greenhouses, 
and these may be used where wild material is not available. In 
early stages where the sporogenous cells have not yet begun to 
round off into spore mother-cells it is sufficient to remove the 
perianth, retaining just enough of the receptacle to hold the sta- 
mens in place. Transverse sections show the six stamens and 
also the young ovary. After the spore mother-cells have begun 
to round off, each stamen should be removed so as to be cut 
separately. It is very desirable to secure stamens showing the 
mitotic figures which occur during the division of the spore 
mother-cell into the four microspores. Since the pollen mother- 
cells are apt to be in approximately the same stage of develop- 
ment throughout the anther, it is worth while to determine 



Spermatophytes 



13 



whether mitotic figures are present before putting the material 
into the fixing agent. The procedure indicated for Phius can be 
followed here. It is not necessary to cut the stamens into 
pieces before fixing, since they are easily penetrated and infil- 
trated ; in later 
stages the sta- 
mens must not be 
cut into pieces, 
since the pollen 
grains are easily 
washed out. 

Transverse 
sections are bet- 
ter for morpho- 
logical purposes; 
but where noth- 
ing is desired ex- 
cept the develop- 
in en t of pollen 
grains from the 
spore mother- 
cells, much more 
material can be 
gotten under a 
cover by using 
longitudinal sec- 
tions. 

Fo r early 

Stages UP tO that * nth f r showing large sporogenous cells, one layer of t; 

D -T deeply shaded in the figure) , and from three to four layers 

^hnwn in Her f)3 Salix petiolaris. X 594. Small portion of transverse section of a nearly 

./<5' KJ> mature anther showing five pollen grains, two tapetal cells, and two layers 

A r\ p | Q f 1 A ' °f wa ^ cells. C, Lilium auratum. X 505. Section of a pollen grain show- 

XX) UeiaiieiQ S \ n g the large tube nucleus and two smaller generative nuclei. Fixed in 

1 , . . chromo-acetic acid and stained in safranin-gentian violet-orange. Z>, 

na^matOXylin IS Lilium tigrinum. X 505. Pollen grain showing tube nucleus in the mid- 

. die and a lenticular cell with the generative nucleus at one end of the grain. 

very satisfactory ; 

in stages like B, C, and D of the same figure, cyanin and eryth- 
rosin often give excellent differentiation, the tube nucleus taking 
the erythrosin, and the generative nucleus the cyanin, while the 




Fig. 63. 

A, Salix tristis. X 594. Transverse section of a portion of a young 

tapetal cells (more 
of wall cells. B, 



132 



Methods in Plant Histology 



starch grains which are often abundant at this stage take a faint 
pink from the erythrosin. However, the safranin-gentian violet- 
orange combination shows nuclear details to better advantage, 
and readily distinguishes the tube and generative nuclei, although 
it does not give such a striking color contrast as the cyanin and 

erythrosin. The starch grains 
are brilliantly stained by the 
gentian violet. 

The mature pollen grain, 
after shedding, may be pre- 
pared by the method already 
described for Pinus. 

Oogenesis. — As in sper- 
matogenesis, the early stages 
will be found in preparations 
of floral development. The 
origin and development of the 
macrospore are easily traced 
in Liliiim. In very young 
stages, before the appearance 
of the integument, the ovary 
may be removed from the 
flower and placed 'directly in 

tic 64. r J 

A , head of Aster. B, pod of Capsella. C, transverse the fixing agent, but in later 
section of ovary of Lilium. The dotted lines show how 

the material should be trimmed before fixing. StagfeS SUCh aS are shown in 

Jigs. 66-71, strips should be cut off from the sides of the ovary 
in order to secure more rapid fixing and more perfect infiltration 
with paraffin. The dotted lines in fig. 64, C, show about how much 
should be cut off. This is a much better plan than to secure 
rapid fixing and infiltration by cutting the ovary into short 
pieces, because the ovules will be in about the same stage of 
development throughout the ovary, and when one finds desirable 
stages like those from which these photomicrographs were taken, 
it is gratifying to have these pieces as long as possible. 

In lilies, and other forms with large cells, the entire series 
shown in figs. 65-70 may be fixed in chromo-acetic acid and 




Spermatophytes 



133 



stained in safranin-gentian violet-orange, but stages earlier than 
that shown in fig. 63 are more satisfactory if stained in Delafield's 
hematoxylin. Stages like that shown in fig. 66 and also those 
between fig. 66 and fig. 67 are the most difficult to stain, and 
only the utmost care and patience will insure first-class prepara- 
tions. The thread on the nucleus in fig. 66 shows a row of 
chromatin granules and soon becomes segmented into twelve 
chromosomes. If the stain is too dense, the thread will probably 
appear smooth throughout all these stages, but if the staining is 
successful, the granules are 
sharply stained by the gen- 
tian violet, while the linin 
thread is stained lightly or 
not at all. In successful 
staining with cyanin and 
erythrosin the granules 
stain blue and the linin red. 
Stages like figs. 6y and 68 
are easily stained, and the 
preparations are exception- 
ally beautiful. Stain for 
twenty-four hours in safra- 
nin (the solution in 50 per 
cent, alcohol is very good) , 
and then rinse in 50 per cent, alcohol until the red color 
disappears from the spindle, but remains bright in the chro- 
mosomes and nucleoli ; stain in gentian violet four to eight 
minutes ; rinse in water about thirty seconds ; stain in aqueous 
orange G fifteen to thirty seconds ; transfer to absolute alcohol, 
and move the slide gently back and forth in order to dehydrate 
as rapidly as possible (three to six seconds will usually be long 
enough); the slide must be taken from the absolute alcohol 
while the gentian violet is still coming out in streams ; treat with 
clove oil ten to thirty seconds, and then drain off the clove oil 
and add a few drops of cedar oil, since the gentian violet fades if 
much clove oil is left in the preparation. If the cedar oil is 




Fig. 65. Lilium philadelphicum. X 710. 

Apex of nucellus contains a large archesporial cell with a 
large nucleus. In Lilium this archesporial cell becomes 
the embryo-sac directly without cutting off any tapetal 
cell or dividing into potential megaspores. Fixed in 
chromo-acetic acid and stained in Delafield's haematoxylin. 



134 



Methods in Plant Histology 



transparent and has a strong odor, do not use it, but remove as 
much of the clove oil as possible. Mount directly in balsam. 
Before transferring to absolute alcohol, a single dip in 95 per 
cent, alcohol may do no damage and is a matter of economy, 
since it avoids carrying so much water into the absolute alcohol. 

When the 
staining is 
properly done, 
the chromo- 
somes will 
show a bright 
red color and 
the spindle a 
brilliantviolet. 
If the action 
of the gentian 
violet be too 
prolonged o r 
poorly ex- 
tracted, the 
chromosomes 
will appear 
violet or red- 
dish violet, 
and such de- 
tails as the re- 
lation of spin- 
dle fibers to 
chromosomes 
will be ob- 
scured. Stages 
shown in figs. 

6q and yo may be found in the same ribbons with stages like figs. 
6j and 68, and are well differentiated by the same treatment. 

Later stages up to fertilization and the first divisions of the 
embryo may be fixed and stained as already described, but 




Fig. 66. Lilium philadelphicum. X 710. 

Nucellus with megaspore. The chromatin thread in the nucleus is very- 
distinct. Fixed in chromo-acetic acid and stained in safranin-gentian violet- 
orange. 



Spermatophytes 



35 



cyanin and erythrosin gives a particularly brilliant effect, 
especially after Carnoy's fluid. At the stage shown in fig. //, 
the male nucleus takes the cyanin, and the oosphere nucleus the 
erythrosin, although at a slightly later stage they stain alike. 





Fig. 67. 



Lilium philadelphicum. X 710. 



Fig. 



First division of the nucleus of the megaspore. (The same nucleus the earlier stages of which are 
shown in figs. 65 and 66.) Fixed in chromo-acetic acid, and stained in safranin- gentian violet-orange. 
Fifteen microns. 

For the later stages in the development of the embryo, Delafield's 
hematoxylin is a far better stain. 

Many of the Composite, like Aster, Taraxacum, Senecio, and 
Silphitim, are excellent for a study of the mature sac and the 
formation of the embryo. The whole head may be cut if trimmed 
as indicated in fig. 64, but for the embryo-sac at the fertilization 
period, and also for the development of the embryo, it is worth 
while to resort to the tedious process of dissecting the ovules 



136 



Methods in Plant Histology 



out from the ovaries. Stages like those shown in figs. 65-jo 
are rather unsatisfactory in Composites. 

Many of the Ranunculaceae are easily studied. Anemone 
patens, var. Nuttalliana, has a very beautiful embryo-sac, the egg, 




Fig. 69. Lilium philadelphicum. X 710. 

Megaspore (embryo-sac) containing two 
daughter-nuclei resulting from the first division of 
the nucleus of the megaspore. A portion of the 
spindle still remains between the two nuclei. 
Fixed in chromo-acetic acid, and stained in safra- 
nin-gentian violet-orange. Fifteen microns thick. 




Fig. 70. 'Lilium philadelphicum. X 710. 

Later stage in the development of the embryo- 
sac. Each of the two nuclei shown in fig. 69 has 
divided. The nuclei are much smaller than those 
in fig. 69. Fixed in chromo-acetic acid, and 
stained in safranin-gentian violet-orange. Fifteen 
microns thick. 



synergids, endosperm nucleus, and antipodals being rather large 
and sharply defined. Hepatica, Caltha, and some species of 
Ranimculus are exceptionally good. 

Development of the Embryo. — The common shepherd's purse 
{Capsella bursa-pastoris) is a favorable form for a study of the 
development of a dicotyl embryo. The stages shown in fig. J2, 



Spermatophytes 



137 



A—F, will be found in pods which are about one-eighth of an 
inch long. These may be put directly into the fixing agent, but 
stages like G and //"are found in pods about three-sixteenths of 
an inch long, and such stages will be more readily fixed, infil- 
trated, and cut if the pods are trimmed, as shown in fig. 64, B, 
before putting them into the fixing 
agent. Cut sections parallel to the 
flat face of the pod. Delafield's 
hematoxylin, without any contrast 
stain, gives the best results which 
we have secured. 

For tracing the development of 
a monocotyl embryo Sagittaria vari- 
abilis can be recommended. Mate- 
rial is abundant, sections are easily 
cut, except in the latest stages, and 
it is not difficult to get a complete 
series. Alisma plantago, which is 
commonly figured in text-books, is 
extremely hard to cut, especially in 
later stages. 

Leaves. — Where only a rapid ex- , 
amination is to be made, free-hand 
sections may be made in great num- 
bers by using" the method employed 
for pine needles. It is easy to get 
good sections of leaves which can FlG - 7I - Lilium p™adeiphicum. 

, . rn . . . . „ Embryo-sac at time of fertilization. A , 

be gotten intO paraltin, but it IS dim- the three antipodals. E, protoplasm of the 

sac. e, polar nuclei fusing to form the 

Cult tO get tender, SUCCulent leaves endosperm nucleus ; the male nucleus is 

about to fuse with the nucleus of the 

intO paraffin Without distortion. Such oosphere. i , the inner integument pt 

r the pollen tube. Fixed in Carnoy s fluid 

lpn V po mPV hp riit in rpllniHin T?r>r and stained in cyanin and erythrosin. Fif- 

leaves may De cut in cenoiain. ror teen microns thick . ( From Baileys c y do- 
a study of the stomata, strip a piece pedia of American Horticulture.) 
of epidermis from the leaf, fix it, stain in Delafield's haematoxylin 
and erythrosin, pass it gradually through the alcohols, clear in 
xylol, and mount in balsam. Lily, tulip, hyacinth, and begonia 
may be suggested as favorable forms. Epidermis from the leaf 




1 3 8 



Methods i?i Plant Histology 



of the common Sedum pupurascens will usually show stomata in 
all stages of development. 

Stems and Roots.- — The earlier stages in the development of 
vascular bundles in stems and roots are well shown in paraffin 




Fig. 72. Capsella bursa-pastoris. X 400. 

A, first division in the embryo cell. B, quadrants. C, octants. D, the dermatogen has been cut 
off. There are eight cells in the suspensor, the lower cell being very large and vesicular. F, differ- 
entiation in plerome and periblem. The plerome cells are shaded. F, the periblem of the root is com- 
pleted at the expense of the upper cell of the suspensor. G, the mitotic figure in the suspensor cell indi- 
cates that the upper suspensor cell by a second contribution is about to complete the dermatogen of the 
root. H, plerome (shaded), periblem, dermatogen (shaded), and first layer of the root cap. Fixed in 
chromo-acetic acid and stained in Delafield's hematoxylin. Ten microns thick. 

sections of young seedlings. The common bean is a favorable 
form, and it is easy to get material. 

Most herbaceous stems and roots, and also the younger 
woody stems and roots, give the best results when cut in celloidin, 
as already described for Pinus. Rumex crispus and Ra?iu?iculus 
repens can be recommended for a study of the vascular bun- 
dles. The cambium is very sharply brought out by Delafield's 



Spermatophytes 



39 



hematoxylin. Petioles or leaf blades of Nuphar imbedded in cel- 
loidin, and stained in safranin and Delafield's haematoxylin, yield 
extremely beautiful preparations, the sclerotic cells taking a 
brilliant red and the cellulose a rich purple. The sections should 
be 20-30 p thick. 

Stems or petioles of the squash or pumpkin are to be pre- 
ferred for demonstrating sieve tubes and companion cells. For 
the more minute de- 
tails of the sieve plate 
it is best to cut out 
small piecesaboutone- 
fourth of an inch long 
and one-eighth of an 
inch square containing 
the vascular bundle. 
These pieces can be 
imbedded in paraffin. 

For demonstrating 
the phellogen and the 
tissue developed from 
it, stems of Geranium 
or Coleus about one- 
fourth of an inch in 
diameter or seedlings 
of Xanthium canadense 
about three-sixteenths 
of an inch in diameter 
can be recommended. 
They can be cut in paraffin, but satisfactory results are more 
uniformly obtained from celloidin sections. 

The stem of Indian corn, imbedded in celloidin and stained 
in safranin and Delafield's haematoxylin, affords a good study of 
monocotyl stem anatomy. 

It must not be forgotten that root tips, besides showing ana- 
tomical structure, furnish ever-ready material for the study of 
karyokinesis. An onion thrown into a pan of water will soon 




Fig. 73. Sparganium eurycarpum. X 53. 

Tranverse section of a root. Delafield's haematoxylin 
and acid fuchsin. Five microns. 



140 Methods i?i Plant Histology 

send out numerous roots. About one-fourth of an inch should 
be cut off from the tip of the root. Fix and stain as directed 
for the mitotic figures in Lilium. While the onion is very avail- 
able material, the figures are not as satisfactory as those to be 
obtained in the root tips of Tradescantia virginica, Iris versicolor, 
Podyphyllum peltatum, Ariscema triphyllum, Cypripedium pubescens, 
and many others. Transverse sections of young roots often show 
a remarkably regular arrangement of the cells, as can be seen in 

fig- 73- 

We are painfully aware that the directions which have been 

given in this series of articles are very incomplete, but it is hoped 
that they will enable the student to devise for himself such 
methods as particular cases may demand. 



CHAPTER XX. 

LABELING AND CATALOGUING PREPARATIONS. 

THE LABEL. 

The labels shown in fig. 62, on p. 126, show as much as 
will generally be found desirable. The date of the collection 
of the material is often needed in addition. The date of making 
the preparation is of no value unless the student is testing the 
permanence of stains or something of that sort. It is hardly 
worth while to write upon the label the names of the stains used, 
for the student will soon learn to recognize the principal stains. 
We should say that the first thing to write upon a label is the 
genus and species of the plant; the next thing would be the 
name of the organ or tissue, and then might be added the date 
of collection : e.g., Marchantia polymorplia, young archegonia, 
April 10, 1 90 1. A hasty sketch on the label will often indicate 
any exceptionally interesting feature in the preparation. To 
facilitate finding such a feature, it is a good plan to mark the 
particular section or sections with ink, the marking being 
always on the underside of the slide so as not to cause any 
inconvenience if an immersion lens should be used. 

CATALOGUING PREPARATIONS. 

As a collection grows, the student will need some device for 
readily locating any particular preparation. Some have their 
slides numbered and catalogued, but all devices of this sort are 
too cumbrous and slow for the practical worker in the labora- 
tory. After several years' experience with a collection which 
now numbers about seven thousand preparations, the following 
method can be confidently recommended : 

Four wooden slide boxes of the usual type will do for a 
beginning ; they should be labeled : Thallophyta, Bryophyta, 
Pteridophyta, and Spermatophyta. As the collection grows 

141 



142 



Methods in Plant Histology 



and new boxes are needed, the classification can be made more 
definite ; e.g., there should be a box labeled Bryophytes Hepaticcz 
and one labeled Bryophytes Musci. As the liverwort collection 
grows, three boxes will be necessary, and should be labeled 
Bryophytes HepaticcB Marchantiales, Bryophytes Hepaticce- Jun- 
germanniales, and Bryophytes //<?/WzV<^ Anthocerotales. It will 
readily be seen that the process can be continued almost indefi- 
nitely, and that new slides may be at any time dropped into their 
proper places. A rather complete label gradually built up in 
this way is shown in Jig. 74: 



BRYOPHYTES 

HEPATIC^ 

Jungermanniales 
Porella platyphyllum 

Archegonia 
Fig. 74- 



CHAPTER XXI. 

A CLASS LIST OF PREPARATIONS. 

Where a regular course in histology is conducted, it is a good 
plan to give each student at the outset a complete list of the 
preparations which he is expected to make. In a three-months' 
course a fairly representative collection of preparations can be 
made. The availability of material determines what a list shall 
be. The following list was recently used by one of the writer's 
classes : 

LIST OF PREPARATIONS. 

THALLOPHYTES. 

ALGJE. 
CYANOPHYCE^. 

i. Wasserbliithe. — The principal forms in this material are : 

(a) Ccelosphceriiim Kutzingiamon. — Colonies in the form of hollow 

spheres. 
(J?) Anabcena gigantea. — Filaments straight. Preparations should show 

vegetative cells, heterocysts, hormogonia, and spores. 
(c) Anabcena flos-aqua. — Filaments curved. Stain on the slide and 

mount in balsam. 

2. Nostoc. — Eosin or erythrosin. Glycerine. 

3. Oscillaria. — Put living material into 10 per cent, glycerine and allow it 
to concentrate. 

4. GloBOtrichia. — Eosin, erythrosin, or iron alum-haematoxylin. Glycerine. 
Should show heterocysts, spores, and vegetative filaments. 

5. Tolypothrix. — Treat like Oscillaria or Gloeotrichia. Should show hetero- 
cysts, hormogonia, and false branching. 

CHLOROPHYCE.E. 

6. Volvox. — Stain some in eosin or erythrosin and some in very dilute 
Delafield's hsematoxylin. Mount in glycerine in a cell deep enough to 
prevent crushing. Try to have antheridia, oogonia, and young colonies 
in each mount. 

7. Vaucheria. — Eosin, erythrosin, or iron alum-haematoxylin. Glycerine. 
Should show antheridia, oogonia, zoospores. 

143 



144 Methods in Plant Histology 

8. Hydrodictyon. — Iron alum-haematoxylin or dilute Delafield's haematoxy- 
lin. Glycerine. Look for young nets inside the older segments. 

9. Spirogyra. — Iron alum-hasmatoxylin. A counter-stain with eosin is a 
great improvement, if successful. Glycerine. Should show chroma- 
tophores, pyrenoids, and nucleus. Eosin, erythrosin, or dilute Delafield's 
haematoxylin will be better for conjugating material. 

1 o. Zygnema. — Same treatment as for Spirogyra to bring out chromatophores, 
pyrenoids, and nucleus. Use iron alum-hgematoxylin for reproductive 
stages. 

11. Diatoms. — Living and fossil forms. Balsam. 

12. Oedogonium. — Use iron alum-hagmatoxylin for oogonia, antheridia, and 
zoospores. Eosin or erythrosin for caps and other vegetative structures. 
Glycerine. 

13. Chara. — Oogonia and antheridia may be mounted whole in a cell deep 
enough to prevent crushing. Glycerine jelly. Paraffin sections of 
oogonia, antheridia, and apical cell. 

ph^:ophyce,e. 

14. Ectocarpus. — Stain some in eosin and some in dilute Delafield's hema- 
toxylin. Glycerine. Each mount should show both unilocular and mul- 
tilocular sporangia. 

15. Fucus vesiculosus. — Antheridial conceptacle with paraphyses and anthe- 
ridia ; oogonial conceptacle with oogonia. Delafield's hematoxylin for 
paraffin sections. Borax carmine for teased preparations. 

RHODOPHYCE.E. 

16. Batrachospermum (or Nemalioii). — Mayer's haem-alum. Glycerine. 
Should show trichogyne, carpogonium, and cystocarp. 

17. Poly sip honia fibrillosa. — Iron alum-haematoxylin or eosin. Glycerine. 
Vegetative structure, tetraspores, cystocarps with carpospores, antheridia. 

FUNGI. 
MYXOMYCETES. 

18. Trichia varia. — Paraffin sections, 5/*. Safranin-gentian violet-orange. 

SCHIZOMYCETES. 

19. Bacteria. — Coccus, Bacillus, and Spirillum forms. Stain on cover-glass 
or slide. 

20. Bacillus anthracis. — In liver of mouse. Paraffin sections, 5 /x. Stain 
in gentian violet, Gram's method. 

PHYCOMYCETES. 

21. Mucor stolonifer. — Stain young sporangia in eosin or dilute Delafield's 
haematoxylin. Zygosporic material may be mounted without staining or 
after a very light staining in dilute Delafield's haematoxylin. Glycerine. 



Class List of Preparations 145 

22. Cystopus candidus on Cakile americana. — Select white blisters which 
have not yet broken open. Paraffin, 5 /x. Safranin-gentian violet-orange. 

23. Cystopus bliti on Amarantus retroflexus. — Cut out small portions of 
leaves in which the oogonia can be seen in abundance. Paraffin, 5 it. 

ASCOMYCETES. 

24. Eurotium. — Eosin or erythrosin. Glycerine. 

25. Pencillium. — Eosin or erythrosin. Glycerine. 

26. Erysiphe commune on Polygonum aviculare. — Strip the fungus from the 
leaf. Paraffin, 5 /*. Safranin-gentian violet-orange. 

27. Uncinula necator on Ampelopsis quinquefolia. — Eosin or erythrosin. 
Mount whole and break the perithecia under the cover. Glycerine or 
balsam. 

28. Xylaria. — Paraffin sections of younger stages. Delafleld's hematoxylin 
and erythrosin. Be sure that some section in each mount shows the 
opening of a perithecium. 

29. Peziza. — Paraffin sections of young apothecia, 5 fi or less ; sections of 
older apothecia, iotcor 15 it. Safranin-gentian violet-orange. 

^CIDIOMYCETES. 

30. Puccinia graminis. — yEcidium stage on barberry leaf. Paraffin. Cyanin 
and erythrosin. Uredospore and teleutospore stage in celloidin or paraffin. 

BASIDIOMYCETES. 

3 1 . Coprinus comatus. — Paraffin. Transverse sections of gills showing trama, 
paraphyses, basidia, and spores. To show the basidium with four spores, 
the sections should be 1 5 n thick. For development of the spores, cut 
5M or less. Safranin-gentian violet-orange. Boletus, Hydnum, and 
Polyporus are treated in the same manner. 

LICHENS. 

32. Parmelia Borreri. — Paraffin, 5 it. Cyanin and erythrosin. 

BRYOPHYTES. 

HEPATIC^. 

33. Ricciocarpus 7iatans. — Paraffin, 10/i or 15 it. Delafleld's haematoxylin. 
Archegonia, antheridia, and sporophytes imbedded in the gametophyte. 

34. Marchantia polymorpha. — Paraffin, 10 it. Archegonia, antheridia, and 
sporophytes. 

35. Pellia epiphylla. — Paraffin, 10/1. Longitudinal sections of sporophyte 
attached to gametophyte. Delafleld's hematoxylin and erythrosin. 

36. Porella platyphyllu??i. — Paraffin, ioai. Delafleld's haematoxylin. Arche- 
gonia, antheridia, sporophyte, and apical cell. 

37. Anthoceros Icevis. — Paraffin, 5 or io/*. Longitudinal and transverse 
sections of sporophyte. Safranin-gentian violet-orange. 



146 Methods i?i Plant Histology 

MUSCI. 

38. Sphagnum. — Leaf buds. Safranin and Delafield's haematoxylin. Tease 
and mount in balsam. 

39. Sphagnum.— Capsule. Paraffin. Delafield's haematoxylin and erythro- 
sin. 

40. Bryum proliferum. — Paraffin. Antheridia, 10/^ ; archegonia, 15 to20ju; 
capsule, 10 At. 

41. Funaria hygro?netrica. — Paraffin. Longitudinal and transverse sections 
of young capsules. Delafield's haematoxylin. 

42. Funaria hygromeirica or any favorable form. Protonema. Place the 
well-cleaned material directly into 10 per cent, glycerine and allow it to 
concentrate. 

PTERIDOPHYTES. 

FILICALES. 

43. Botrychiwn virginianum. — Paraffin. Stain rhizome, stipe, and root in 
safranin and Delafield's haematoxylin. Stain sporangia in Delafield's 
hematoxylin. 

44. Isoetes echinospora. — Transverse section of stem. Celloidin. Safranin 
and Delafield's hasmatoxylin. 

45. Isoetes echinospora. — Paraffin. Longitudinal sections of microsporangia 
and megasporangia. Safranin-gentian violet-orange. 

46. Pteris aquilina. — Free-hand sections. Safranin and Delafield's haematox- 
ylin, or iodine green and fuchsin. Balsam. 

47. Pteris cretica. — Paraffin. Transverse sections of leaves with sporangia. 
Delafield's haematoxylin and erythrosin or safranin-gentian violet-orange. 

48. Adia?itum cuneatum.— Prothallia mounted whole in glycerine. Paraffin 
sections of antheridia and archegonia. 

49. Marsilea quadrifolia. — Paraffin. Longitudinal and transverse sections 
of sori. Safranin-gentian violet-orange. 

50. Azolla carolinense. — Paraffin. Vertical sections of the whole plant 
showing micro- and megasporangia. 

EQUISETALES. 

51. Equisetum arvense. — Prothallia in glycerine. Stem tips in paraffin. 
Transverse section of stem in celloidin. 

LYCOPODIALES. 

52. Lycopodiuni lucidulum. — Transverse section of stem. Free-hand or in 
celloidin. Safranin and Delafield's hasmatoxylin. 

53. Lycopodiuni inundatum. — Paraffin. Longitudinal sections of strobilus. 

54. Selagi?iella. — Paraffin. Longitudinal sections of rather mature strobili. 
Cyanin and erythrosin. 



Class List of Preparations 1 4 7 

SPERMATOPHYTES. 

GYMNOSPERMS. 

55. Pinus Laricio. — Transverse section of needles and young stem. Free- 
hand or celloidin. Safranin and Delafield's hematoxylin. 

56. Pinus strobus. — Free-hand sections of well-seasoned wood. Iodine 
green and fuchsin, or safranin and Delafield's hematoxylin. 

57. Pinus Laricio. — Paraffin. Ovule with archegonia. Safranin-gentian 
violet-orange. 

58. Pinus sylvestris or P. Laricio. — Paraffin. Embryos. 

59. Pi?ius Laricio. — Paraffin. Longitudinal section of mature staminate 
strobilus. Safranin-gentian violet-orange. 

ANGIOSPERMS. 
MONOCOTYLS. 

60. Lilium philadelphicum. — Paraffin, 10 ft. Transverse sections of ovary ; 
transverse sections of anthers. 

61. Tradescantia virginica. — Paraffin, 5 and 10 fx. Longitudinal sections of 
root. Stain for mitosis. 

62. Alliimi. — Paraffin. Transverse sections of older roots for vascular 
system. 

63. Lilium longiflorum. — Strip epidermis from leaf and stain for stomata. 
Cut sections of leaf in paraffin, 10 to 20 /*. 

64. Zea mat's. — Celloidin. Longitudinal and transverse sections of stem. 

DICOTYLS. 

65. Nuphar advena. — Celloidin. Section of leaf 15 to 40^ thick. Safranin 
and Delafield's hematoxylin. 

66. Capsella bursa-pastoris. — Paraffin. Floral development, 5 /x. Embryos, 
5 to 10 /x. Stain both in Delafield's hematoxylin without any contrast 
stain. 

67. Taraxacum officitiale. — Paraffin. Floral development, 5 /*. Embryo- 
sac, IO tO 15 fi. 

68. Aiiemone patens. — Paraffin. Embryo-sac. 

69. Smilax herbacea. — 'Celloidin. Transverse section of root. Safranin 
and Delafield's hematoxylin. 

70. Xanthittm ca?iadense. — Celloidin. Transverse section of stem of seed- 
ling to show cambium and phellogen. 

71. Tilia americana. — Celloidin or free-hand. Transverse sections of small 
stems one-eighth to one-fourth of an inch in diameter. Safranin and 
Delafield's hematoxylin. 

In making the mounts the order indicated in the list should 
not be followed. Begin with free-hand sections, then study the 



148 Methods in Plant Histology 

glycerine method, and then devote a large portion of the time to 
the paraffin method. Let the celloidin method come last. 

It is neither possible nor desirable that each student should 
in every case go through all the processes from collecting mate- 
rial to labeling. Some of the material may be in 70 per cent, 
alcohol, some in formalin, some in glycerine, and some in paraffin. 
One student may imbed in paraffin enough of the A7iemone for 
the whole class, another may imbed the Lilium stamens, and by 
such a division of labor a great variety of preparations may be 
secured without a corresponding demand upon the time of the 
individual. 



CHAPTER XXII. 

FORMULA FOR REAGENTS. 

FIXING AGENTS. 
Carnoy's Fluid. — 

Absolute alcohol, 6 parts. 
Chloroform, 3 parts. 
Glacial acetic acid, 1 part. 
Strong Chromo-Acetic Solution. — 

Chromic acid, 1 g. 
Glacial acetic acid, 1 cc. 
Water, 98 cc. 
Weak Chromo-Acetic Solution (Schaffner's) . — 

Chromic acid, 0.3 g. 
Glacial acetic acid, 0.7 cc. 
Water, 99 cc. 

Medium Chromo-Acetic Solution. — 

Chromic acid, 0.7 g. 
Glacial acetic acid, 0.5 cc. 
Water, 100 cc. 
Flemming's Fluid (weaker solution). — 
( 1 per cent, chromic acid, 25 cc. 
A < 1 per cent, acetic acid, 10 cc. 

( Water, 55 cc. 
B. 1 per cent, osmic acid, 55 cc. 

Keep the mixture A made up, and add] B as Jthe [reagent is 
needed for use, since it does not keep well. 

Flemming's Fluid (stronger solution). — 

1 per cent, chromic acid, 45 cc. 

2 per cent, osmic acid, 12 cc. 
Glacial acetic acid, 3 cc. 

Merkel's Fluid.— 

1.4 per cent, solution of chromic acid, 25 cc. 
1.4 per cent, solution of platinic chloride, 25 cc. 

Hermann's Fluid. — 

1 per cent, platinic chloride, 15 parts. 
Glacial acetic acid, 1 part. 

2 per cent, osmic aid, 4 or 2 parts. 

149 



I 5 o Methods in Plant Histology 

Picric Acid. — 

Picric acid, i g. 

Water, or 70 per cent, alcohol, 100 cc. 

Corrosive Sublimate. — 

Bichloride of mercury, 4 g. 

Glacial acetic acid, 2 cc. 

Water, or 70 per cent, alcohol, 100 cc. 

Formalin (weaker solution). — 
Formalin, 2 cc. 
Water, 98 cc. 

Formalin (stronger solution).— 
Formalin, 4 cc. 
Water, 100 cc. 

Osmic Acid. — 

Osmic acid, 1 cc. 
Distilled water, 100 cc. 

The bottle in which the solution is to be kept, and also the 

glass tube in which the acid is sold, must be thoroughly cleaned. 

Break off the end of the tube, and drop both tube and acid into 

the distilled water. 

STAINS. 

Delafield's Hematoxylin. — "To 100 cc. of a saturated solution 
of ammonia alum add, drop by drop, a solution of I g. of hema- 
toxylin dissolved in 6 cc. of absolute alcohol. Expose to air 
and light for one week. Filter. Add 25 cc. of glycerine and 
25 cc. of methyl alcohol. Allow to stand until the color is suf- 
ficiently dark. Filter and keep in a tightly stoppered bottle." 
(Stirling and Lee.) 

The solution should stand for at least two months before it 
is ready for using. 

Erlich's Hematoxylin. — 

Distilled water, 50 cc. 
Absolute alcohol, 50 cc. 
Glycerine, 50 cc. 
Glacial acetic acid, 5 cc. 
Haematoxylin, 1 g. 
Alum in excess. 

Keep it in a dark place until the color becomes a deep red. 

If well stoppered, it will keep indefinitely. 



Formula for Reagents 151 



Boehmer's Hematoxylin. — 

Hematoxylin, 1 g. 



Absolute alcohol, 12 cc. 

B j Alum > 1 g- 

J Distilled water, 240 cc. 

The solution A must ripen for two months. When wanted 
for use, add about 10 drops of A to 10 cc. of B. Stain ten to 
twenty minutes. Wash in water and proceed as usual. 

Mayer's Haem-Alum. — Hematoxylin, 1 g., dissolved with 
heat in 50 cc. of 95 per cent, alcohol and added to a solution of 
50 g. of alum in a liter of distilled water. Allow the mixture to 
cool and settle ; filter ; add a crystal of thymol to preserve from 
mold. (Lee.) 

It is ready for use as soon as made up. Unless attacked by 
mold, it keeps indefinitely. 

Haidenhain's Iron Alum-Haematoxylin. — This stain was intro- 
duced by Haidenhain in 1892 and has gained a well-deserved 
popularity with those engaged in cytological work. Two solu- 
tions are used, and they are never mixed : 

A. One and one-half to 4 per cent, aqueous solution of 
ammonia sulphate of iron. (At present we use a 3 per cent. 
solution.) 

B. One-half per cent, aqueous solution of hematoxylin. 
Greenacher's Borax Carmine. — 

Carmine, 3 g. 

Borax, 4 g. 

Distilled water, 100 cc. 

Dissolve the borax in water and add the carmine, which is 
quickly dissolved with the aid of gentle heat. Add 100 cc. of 
70 per cent, alcohol and filter. (Stirling.) 

Alum Carmine. — A 4 per cent, aqueous solution of ammonia 
alum is boiled twenty minutes with 1 per cent, of powdered car- 
mine. Filter after it cools. (Lee.) 

Alum Cochineal. — 

Powdered cochineal, 50 g. 

Alum, 5 g. 

Distilled water, 500 cc. 



152 Methods in Plant Histology 

Dissolve the alum in water, add the cochineal, and boil ; 
evaporate down to two-thirds of the original volume, and filter. 
Add a few drops of carbolic acid to prevent mold. (Stirling.) 

Picro-Carmine. — 

Picro-carmine (picro-carminate of ammonia), 1 g. 
Water, 100 cc. 
Eosin. — 

Eosin, 1 g. 

Water, or 70 per cent, alcohol, 100 cc. 

General Formula for Anilins. — Make a 3 per cent, solution of 
anilin oil in distilled water ; shake well and frequently for a day; 
add enough alcohol to make the whole mixture about 20 per 
cent, alcohol ; add 1 g. of cyanin, erythrosin, safranin, gentian 
violet, etc., to each 100 cc. of this solution. 

Iodine Green. — 

Iodine green, 1 g. 

70 per cent, alcohol, 100 cc. 

Methyl Green. — 

Methyl green, 1 g. 
Glacial acetic acid, 1 cc. 
Water, 100 cc. 

If the preparation is to be mounted in balsam, a slight trace 
of acetic acid and also a trace of methyl green should be added 
to the absolute alcohol used for dehydrating. 

Fuchsin. — 

Fuchsin, 1 g. 

95 per cent, alcohol, 100 cc. 

Water, 100 cc. 

ZiehPs Carbol Fuchsin. — 

Fuchsin, 1 g. 

Carbolic-acid crystals, 5 g. 
95 per cent alcohol, 10 cc. 
Water, 100 cc. 

Fuchsin and Iodine Green Mixtures. — Two solutions are kept 
separate, since they do not retain their efficiency long after they 

are mixed. 

o. 1 g. fuchsin (acid). 
50 cc. distilled water. 



Formula for Reagents I 5 3 

j 0.1 g. iodine green. 
( 50 cc. distilled water. 
(100 cc. absolute alcohol. 
C I 1 cc. glacial acetic acid. 
( 0.1 g. iodine. 

Stain in equal parts of A and B. Transfer from the stain 
directly to solution C and from C to xylol. 

Another Formula. — 

Acid fuchsin, 0.5 g. 

Water, 100 cc. 
j Iodine green, 0.5 g. 
( Water, 100 cc. 

Mix a pipette full of A with a pipette full of B ; stain two to 
eight minutes ; dehydrate rapidly and mount in balsam. 

Safranin. — 

Safranin, 1 g. 

95 per cent, alcohol, 50 cc. 

Water, 50 cc. 

Gentian Violet. — 

Gentian violet, 1 g. 
95 per cent, alcohol, 20 cc. 
Water, 80 cc. 
Anilin, 3 cc. 
Orange G. — 

Grange G, 1 g. 
Water, 100 cc. 

Bismark Brown. — 

Bismark brown, 2 g. 
70 per cent, alcohol, 100 cc. 
Nigrosin. — 

Nigrosin, 1 g. 
Water, 100 cc. 

Grants Solution. — 

Iodine, 1 g. 

Iodide of potassium, 2 g. 

Water, 300 cc. 



INDEX 



INDEX. 



The references are to pages. 

Acid alcohol, formulas, 35, 43. 

-#Lcidiomycetes, 84. 

Air bubbles, 32 

Alcohol, for fixing, 25 ; formulae, 9. 

Algae, 63. 

Alisma, embryo, 137. 

Alum carmine, 40. 

Alum cochineal, 40. 

Ammonia, 35. 

Anabaena, 63. 

Angiosperms, embryo, 136, 138; fertiliza 
tion, 135, 137; floral development, 129 
oogenesis, 132 ; root, 139 ; spermato 
genesis, 130, 131 ; stem, 138. 

Anilins, formulae, 41, 152. 

Antheridia, Chara, 72, 73; ferns, 105 
joy ; liverworts, gi; mosses, 11, gy. 

Anthers, fixing, 31 ; staining, 131, 132. 

Anthoceros, g4. 

Anthrax, yy. 

Apparatus, I. 

Archegonia, ferns, 106, 107 ; pines, 121 
122, 123 ; liverworts, g2; mosses, gy, 98 

Arisaema, root tip, 140. 

Ascomycetes, 80. 

Aspidium, sporangia, 108. 

Aster, 18, 132. 

Asterella, 89, gi. 

Azolla, fixing, 31 ; infiltrating, 112. 

Bacteria, yy. 

Balsam, mounting in, 22. 

Basidiomycetes, 87. 

Batrachospermum, yj. 

Beggiatoa, 78. 

Bismark brown, 45. 

Boletus, 88. 

Borax carmine, 40. 

Botrychium, anatomy, 109 ; prothallia 

104; sporangia, 108. 
Bryophytes, 89. 
Bryum, antheridia, 97 ; capsule, gg. 



Italic figures indicate illustrations. 

Capsella, floral development, 129; pod, 
132 ; embryo, 138. 

Carmines, 39. 

Carnoy's fluid, formula, 26. 

Cataloguing preparations, 141, 142. 

Cedar oil, 43. 

Celloidin, 15 ; method, 55. 

Centrosomes, 39. 

Chara, paraffin sections, 72, 73; glycerine 
mounts, 58. 

Chlorophyceae, 65. 
; Chorda, 75. 

Chromosomes, 39. 

Chromic acid group, 26. 

Chromo-acetic acid, formulae, 28. 
' Cladophora, 65, 67. 

Clearing, 16, 22; clearing agents, 10. 

Clove oil, 41. 

Cnicus, floral development, 130. 

Ccelosphaerium, 63. 

Coleus, stem anatomy, 139. 
' Collema, 88. 

Conocephalus, 89, 94. 

Coprinus, 87. 

Corn, stem anatomy, 139. 

Corrosive sublimate, 30. 

Cover-glasses, cleaning, 22. 

Crucibulum, 88. 

Cutting paraffin, 19; celloidin, 56. 

Cyanin, formula, 8. 

Cyanin and erythrosin, staining in, 42. 

Gyanophyceae, 63. 

Cyathus, 88. 
. Cypripedium, root tip, 140. 

Cyrtomium, sporangia, 108. 

Cystopus, yg. 

Diatoms, 69, yo. 
Dehydrating, 14. 
Desmids, 71, y2. 
Dichonema, 88. 

157 



i 5 8 



Methods in Plant Histology 



Ectocarpus, 73. 

Eosin, formula, 9. 

Equisetum, 115, 116. 

Erysiphe, 83. 

Erythrosin, formula, 8 ; staining in, 42. 

Eurotium, 81. 

Filicinese, 103. 

Fixative, Mayer's albumin, 20. 

Fixing, general hints, 31. 

Flemming's fluid, formula, 28. 

Floral development, 129. 

Formalin, 31. 

Formula? for reagents, 149. 

Fuchsin acid, formula, 8 ; staining in, 44. 

Fucus, 74. 

Fuligo, 78. 

Funaria, antheridia, 97; capsule, q8, qq. 

Fungi, 77. 

Gentian violet, 8, 43. 
Geranium, stem anatomy, 139. 
Glceeotrichia, 65. 
Glycerine jelly, 58. 
Glycerine method, 58. 
Gymnosperms, 119. 

Hsem-alum, Mayer's, 8, 38. 

Hematoxylin, Boehmer's, 38 ; Dela- 
field's, 7, 35 ; Erlich's, 37; Haiden- 
hain's iron alum, 8, 38 ; Kleinenberg's, 
37. 

Hardening, 14. 

Hemitrichia, 78. 

Hepaticae, 89. 

Hermann's fluid, formula, 29. 

Hydnum, 88. 

Hydrodictyon, 66, 67. 

Hypoxylon, 83. 

Imbedding in paraffin, 18; in celloidin, 

55- 
Iodine green, formula, 8 ; staining in, 44. 
Iris, root tip, 140. 
Isoetes, 117. 

Karyokinesis, 39; in Pinus, 1 19, 122, 123; 
in Lilium, 135, 136; in Osmunda, no; 
in root tips, 139. 



Killing and fixing, 13, 25. 

Labels, 126, 141. 

Laminaria, 75. 

Leptothrix, 78. 

Lichens, 88. 

Lilium, archesporium, 133; fertilization, 
137; fixing anthers, 31; fixing ovaries, 
132; germination of megaspore, 13s, 
136; megaspore, 134. 

List of preparations, 143. 

Lycoperdon, 88. 

Lycopodinese, 117. 

Lycopodium, 118. 

Marchantia, 89, 90, 91, Q2, 94. 

Marsilea, antheridium, 112; archegonium, 

in; imbedding megaspores, 19, 112; 

procuring material, III. 
Merkel's fluid, formula, 29. 
Mica slips, 65. 
Microscope, I, 2. 
Microsphaera, 82. 
Microtome, hand, /; sliding, 2. 
Mildews, 82. 
Mosses, 97. 
Mucor, 7Q. 
Myxomycetes, 78. 

Nigrosin, 45. 
Nostoc, 65. 
Nummularia, 83. 
Nuphar, sclerotic cells, 139. 

Oedogonium, 72. 
Orange G, formula, 8. 
Oscillaria, 64. 
Osmotic apparatus, 15. 
Osmunda, prothallia, 103; sporangia, 
no. 

Paraffin bath, 4; infiltration, 17; removal 
of, 21; summary of method, 23. 

Parmelia, 88. 

Pellia, antheridia, 91 ; sporophyte, Q3 ; 
spores, 94. 

Peltigera, 88. 

Penicillium, 81. 

Peronospora, 81. 



Index 



159 



Peziza, 84. 

Phaeophyceae, 73. 

Phycomycetes, 79. 

Physcia, 88. 

Picric acid, 14, 30. 

Pinus, embryo, 121 ; leaves, 124 ; oogen- 
esis, 121; spermatogenesis, 119, 120; 
stems and roots, 124, 125 ; ventral ca- 
nal cell, 123. 

Podophyllum, root tip, 140. 

Pollen, of Lilium, 131 ; of Pinus, 120. 

Polyporus, 88. 

Polysiphonia, 76. 

Polytrichum, antheridia, 97. 

Prothallia, 1 1, 26, 103, /05, in, 112. 

Protonema, 11, 101. 

Pteridophytes, 103. 

Pteris, antheridia, 107 ; archegonia, 106; 
prothallia, 103, /05; rhizome, no; 
sporangia, 108. 

Ptilidium, go. 

Puccinia, 84, 8j. 

Pythium, 81. 

Ranunculus, 138. 

Razor, 4. 

Reagents, 7. 

Rhodophyceae, 75. 

Riccia, 89, 90. 

Ricciocarpus, 89, <?j. 

Rivularia, 64. 

Roots, of seed plants, 138, zjg; of Equi- 

setum, 116; tips for karyokinesis, 139, 

140. 
Rumex, 138. 

Saccharomyces, 80. 

Safranin, formulae, 8, 42; safranin-gentian 

violet-orange method, 42. 
Sagittaria, embryo, 137. 
Salix, 130, 131. 
Saprolegnia, 81. 
Schizomycetes, 77. 
Scytonema, 65. 
Selaginella, 113, 117. 
Silphium, ovules, 119. 
Spermatophytes, 119. 



Sphagnum, 100. 

Spirillum, 77. 

Spirogyra, 13; fixing, 26, 67, 68; imbed- 
ding, 69. 

Sporangia, of ferns, 108. 

Staining, 33 ; analytical value, 49 ; Fi- 
scher's experiments, 50; general re- 
mark, 47; practical hints, 53; Stras- 
burger's theory, 48. 

Stains, formulae, 7. 

Staphylococcus, 77. 

Starch, 11. 

Stemonitis, 78. 

Stem, of seed plants, 138. 

Stender dish, j. 

Stereum, 88. 

Sticta, 88. 

Taraxacum, 129. 
Teleutospores, 85, 86. 
Temporary mounts, 11. 
Thamnidium, 81. 
Tolypothrix, 65. 
Tradescantia, root tips, 140 
Tremella, 88. 
Trichia, 78. 
Turn table, j. 

Uncinula, 82. 
Uredospores, 85, 86. 
Ustilago, 86. 
Ustilina, 83. 
Usnea, 88. 

Vaucheria, 27, 66. 
Volvox, 65. 

Wasserbluthe, 63. 

Washing, 14. 

Webbera, archegonia, 97. 

Xanthium, stem anatomy, 139. 

Xylaria, 83. 

Xylol, removal of, 21. 

Zea, stem anatomy, 139. 
Zygnema, 6g. 



Jmly-e 1901 



LIBRARY OF CONGRESS 



III 






Mini mil urn mm mi! Hill 
005 386 179 9 



